Handbook of Neurochemistry and Molecular Neurobiology: Neural Signaling Mechanisms [3 ed.] 9780387303383, 9780387303703

Table of contents :
Front Matter....Pages i-xv
Back Matter....Pages 1-13
....Pages 15-25

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Signaling in Development

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Calcium Signaling and Cell Fate Determination During Neural Induction in Amphibian Embryos

M. Moreau . S. E. Webb . I. Ne´ant . A. L. Miller . C. Leclerc

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4

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Calcium Is Involved in the Choice Between Neural and Epidermal Fate . . . . . . . . . . . . . . . . . . . . . . . . 4

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Dihydropyridine-Sensitive Ca2+ Channels and Neural Induction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

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Imaging Ca2+ Transients During Neural Induction in Intact Amphibian Embryos . . . . . . . . . . . . . . . Ca2+ Signaling Activity Begins at the Blastula Stage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ca2+ Activity Was Found to be Restricted Exclusively to the Dorsal Ectoderm . . . . . . . . . . . . . . . . . . . . . In Intact Embryos, Inhibition of Calcium Signaling Disrupts Neural Induction . . . . . . . . . . . . . . . . . . .

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5 Are There Ca2+-Sensitive Target Genes? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 5.1 Identification of New Ca2+ Target Genes Involved in Neural Induction . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 6 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 6.1 How Gene Expression During Neural Induction Might Be Controlled by Ca2+ . . . . . . . . . . . . . . . . . . . . 10

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2009 Springer ScienceþBusiness Media, LLC.

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Calcium signaling and cell fate determination during neural induction in amphibian embryos

Abstract: Experiments performed with isolated ectoderm explants from Xenopus laevis embryos suggest that neural determination is a ‘‘by default’’ mechanism, which occurs when bone morphogenetic proteins (BMPs) are antagonized by extracellular antagonists. BMPs are responsible for the determination of epidermis. However, Ca2+ imaging of intact Xenopus embryos reveals patterns of Ca2+ transients that are generated via the activation of dihydropyridine (DHP)-sensitive Ca2+ channels in the dorsal ectoderm but not in the ventral ectoderm. These increases in the concentration of intracellular Ca2+ ([Ca2+]i) appear to be necessary and sufficient to direct the ectodermal cells toward a neural fate as increasing the [Ca2+]i artificially results in neuralization of the ectoderm. The construction of a subtractive cDNA library between untreated and caffeine-treated (i.e., to increase [Ca2+]i) ectoderms led to the identification of early Ca2+-sensitive target genes expressed in the neural territories. One of these genes, which encodes an arginine methyl transferase, was found to control the expression of the early proneural gene, Zic3. Here, we discuss the possibility of an alternative model to the current ‘‘by default’’ mechanism, where Ca2+ plays a central regulatory role, and epidermal determination only occurs when the Ca2+-dependent signaling pathways are inactive. List of Abbreviations: BCNE, Blastula Chordin and Noggin-expressing center; BMP, bone morphogenetic proteins; DHP, dihydropyridine; DRE, Downstream Regulatory Element; DREAM, DRE antagonist modulator; FGF, fibroblast growth factors; TRPs, transient receptor potential channels

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Introduction

In amphibian embryogenesis, the formation of the nervous system occurs during gastrulation as a result of a process called neural induction. In the last 15 years, it has been suggested that neural induction results from dorsalizing signals such as noggin, chordin, follistatin, XnR3, and cerberus (which originate in the dorsal mesoderm), opposing the normal action of ventralizing signals such as the bone morphogenetic proteins (BMPs), which are derived from the ectoderm and are responsible for the determination of the ectoderm (reviewed by Sasai and De Robertis, 1997). There is mounting evidence to suggest, however, that antagonizing BMP signaling alone is not sufficient to promote neural induction and that other signaling components such as fibroblast growth factors (FGFs) are also required (Delaune et al., 2005; Stern, 2005). In this chapter, we suggest that an additional signaling pathway may also contribute to neural induction where transient increases in the concentration of intracellular calcium ([Ca2+])i play a key role in controling the binary determination decision (i.e., epidermal versus neural tissue). As a result, we present a new model to include the role of Ca2+ in neural induction and to reevaluate the concept of ‘‘by default’’ neural induction.

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Calcium Is Involved in the Choice Between Neural and Epidermal Fate

Ectoderm (i.e., derived from the animal cap) removed at the blastula stage is known to exhibit a high level of morphogenetic plasticity. Without inducing factors it develops into atypical epidermis and with appropriate neural inducers such as noggin, these cells express a variety of neural markers, thus indicating a neural fate (Okabayashi and Asashima, 2003). Barth and Barth (1964) were the first to suggest that in Rana pipiens embryos, Ca2+ is required to trigger neuralization. Following on from this early report, it was shown that the dissociation of Xenopus laevis and Pleurodeles waltl animal caps in Ca2+- and Mg2+-free medium directed cells toward a neural fate (Grunz and Tacke, 1989; Saint-Jeannet et al., 1989, 1990, 1993). More recently, Leclerc et al. (2001) demonstrated that the dissociation of Xenopus laevis animal caps in Ca2+-free medium triggers an increase in [Ca2+]i. This increase was shown to be because of a release of Ca2+ from internal stores, resulting from the inversion of the gradient of concentration in Ca2+ between intra- and extracellular compartments (Leclerc et al., 2001). In addition, neuralization by dissociation was blocked when the animal cap cells were preloaded with the Ca2+ chelator, BAPTA, as suggested by the observation that the neural marker NCAM was not expressed (Leclerc et al., 2001). These results suggest that in Xenopus (and other amphibian) animal caps a Ca2+-dependent signal is necessary both to trigger neuralization of the ectoderm and to inhibit epidermal determination.

Calcium signaling and cell fate determination during neural induction in amphibian embryos

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Dihydropyridine-Sensitive Ca2+ Channels and Neural Induction

Early work on neural induction in amphibians identified the lectin concanavalin A (Con A) as a potent neural inducer when added to animal caps of both anurans and urodeles (Takata et al., 1981; Gualandris et al., 1983, 1985). Indeed, it is still the only lectin that has been shown to have inducing activity. In addition, ConA has been shown to bind to Ca2+ channels and trigger their activation (Greenberg et al., 1987). More recently, it was shown that an increase in Ca2+ occurs during ConA-stimulated neural induction in Pleurodeles waltl animal caps (> Figure 1-1a) and that the use of dihydropyridine (DHP)sensitive Ca2+ channel antagonists inhibit this ConA-induced neural induction (Moreau et al., 1994). Addition of noggin to animal caps also triggers an increase in [Ca2+]i (> Figure 1-1b). This increase has a duration of 10–20 min and represents about a 20% increase in the resting level of [Ca2+]i (Moreau et al., 1994; Batut et al., 2005). Antagonists of DHP-sensitive Ca2+ channels, such as nifedipine or nimodipine, completely block this increase in [Ca2+]i (> Figure 1-1b) and also inhibit neural induction (Leclerc et al., 1997). On the other hand, animal caps that are treated with specific agonists of DHP-sensitive Ca2+ channels, such as S(-)Bay K 8644, generate a transient increase in [Ca2+]i with a duration of 20 min (> Figure 1-1c). This increase is sufficient, even in an active BMP context, to trigger not only the expression of neural markers but also the formation of neurons and glial cells (Moreau et al., 1994). In addition, methylxanthines, such as caffeine or theophyline, which are known to stimulate the release of Ca2+ from internal stores (> Figure 1-1c), are also potent neural inducers (Moreau et al., 1994; Leclerc et al., 1995). These latter experiments suggest that no matter which store it is released into the cytosol from (i.e., either intracellular or extracellular), Ca2+ plays a crucial role as a trigger to neuralize the ectoderm.

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Imaging Ca2+ Transients During Neural Induction in Intact Amphibian Embryos

4.1 Ca2+ Signaling Activity Begins at the Blastula Stage The role of Ca2+ in neural induction has also been examined in intact Xenopus embryos in vivo. Using the Ca2+-sensitive photoprotein, aequorin, in conjunction with a custom-designed photon-imaging microscope (Webb et al., 1997), the Ca2+ dynamics that occur in Xenopus ectodermal cells were directly visualized. Ca2+ signaling activity begins at the blastula stage (i.e., stage 8), long before the onset of gastrulation (i.e., before mesoderm invagination), where the Ca2+ transients were found to be localized in the most anterior part of the dorsal ectoderm (Leclerc et al., 2000). These observations suggest that neural induction might be initiated earlier than was previously thought, but this possibility needs further investigation. Recent studies have demonstrated that neural induction requires the combined activity of the Nieuwkoop Center and the Blastula Chordin and Noggin-expressing center (BCNE) located in dorsal animal cells (Kuroda et al., 2004). The BCNE contains the prospective neuroectoderm and Spemann organizer precursor cells, and is required for brain formation. We propose that the Ca2+ transients observed in the dorsal ectoderm during the blastula stage might be localized in the BCNE. Thus, these Ca2+ transients might prove to be the first directly visualized events linked to neural induction.

4.2 Ca2+ Activity Was Found to be Restricted Exclusively to the Dorsal Ectoderm As gastrulation proceeds, the number and intensity of the Ca2+ transients increase, to reach a peak of activity by midgastrulation (i.e., stage 11–11.5). This activity was found to be restricted exclusively to the dorsal ectoderm (i.e., the tissue where neural induction takes place) and never occurred in the ventral ectoderm cells, (i.e., which do not receive neural inductive signals; Leclerc et al., 2000). > Figure 1-2a shows an example of a series of localized Ca2+ transients that occurred in the dorsal ectoderm of a representative intact embryo between stages 8 and 13.

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Calcium signaling and cell fate determination during neural induction in amphibian embryos

. Figure 1-1 Photometric measurements of the fluorescence changes (F/F0) of the Ca2+ indicator Fluo-3 in (a) Pleurodeles waltl and (b, c) Xenopus laevis animal caps on treatment with (a) ConA and Succinyl-ConA (S-ConA); (b) noggin, both alone and with nimodipine; and (c) caffeine and S(-)BayK. (a) ConA stimulates an increase in [Ca2+]i in competent (blastula) animal caps, whereas S-ConA does not affect the [Ca2+]i. (b) Noggin stimulates an increase in [Ca2+]i in animal caps via dihydropyridine (DHP)-sensitive Ca2+ channels; when noggin is applied to animal caps that are pretreated with nimodipine, a specific antagonist of DHP-sensitive Ca2+ channels, the normal noggin-induced [Ca2+]I increase is blocked. (c) Treatment with caffeine, which releases Ca2+ from internal stores, or the DHP-agonist S() BayK 8644 also stimulates an increase in [Ca2+]i in animal caps

In an attempt to simplify the experimental model, Keller open-face explants (Keller and Danilchik, 1988) have also been used, as a two-dimensional system to study neural induction (> Figure 1-3). It has been reported that this two-dimensional explant system is sufficient to reproduce many aspects of neural induction observed in vivo, such as the expression of neural marker genes, neuronal differentiation, and the

Calcium signaling and cell fate determination during neural induction in amphibian embryos

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. Figure 1-2 Examples of the Ca2+ transients that occur in the dorsal ectoderm of (A) intact embryos and (B) open-face explants during neural induction. (A) Images of a representative embryo at stages 8-9, 10, 11, 12 and 13, on to which were superimposed small circles to mark the positions of the localized Ca2+ transients observed during each stage. AP, VP, D, BPL, and BP are animal pole, vegetal pole, dorsal, blastopore lip and blastopore, respectively. (B) An example of the Ca2+ signals (highlighted by open circles) observed in a representative open-face explant over a period of ~4.5 h. The explant was prepared at ~9 hpf and imaging started at ~11 hpf. Each panel is an aequorin-generated image (which represents 120 s of accumulated luminescence) that was superimposed on the corresponding bright-field image. The number in the lower right of each panel represents the time elapsed since the generation of the first Ca2+ transient (taken as 0 h). Ect. and Mes. are ectoderm and mesoderm, respectively, and the dotted line in panel “a” shows the boundary between the two. Scale bars are (A) 500 mm and (B) 200 mm

induction of a regionalized neural plate along the antero-posterior axis (Doniach, 1992). The Ca2+ imaging data also support this general finding, as a series of localized Ca2+ transients were observed to start from the most anterior part of the open-face explant (> Figure 1-2B; Leclerc et al., 2003), a signaling phenomena similar to that observed in the intact embryo (> Figure 1-2A; Leclerc et al., 2000).

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Calcium signaling and cell fate determination during neural induction in amphibian embryos

. Figure 1-3 Sagittal section of a Xenopus embryo at stage 10 (9 hpf). Open-face explants (dotted box) are prepared by excising tissue (both ectoderm and mesoderm) from the lip of the blastopore up to the animal pole after which it is cultured flattened under a coverslip. The ectoderm consists of the animal cap (AC) and noninvoluting marginal zone (NIMZ), and the mesoderm consists of the involuting marginal zone (IMZ). In this explant system, neural inducing signals can only travel in a planar pathway from the mesoderm to the ectoderm, as vertical signals from the involuting mesoderm are effectively eliminated. AP, VP, D, and V are animal pole, vegetal pole, dorsal, and ventral, respectively. Ect. and Mes. are ectoderm and mesoderm, respectively. BP is blastopore

4.3 In Intact Embryos, Inhibition of Calcium Signaling Disrupts Neural Induction Furthermore, intact Xenopus embryos and explants also yielded similar results on treatment with either the Ca2+ chelator BAPTA or specific antagonists of the DHP-sensitive Ca2+ channels, where neural induction was significantly disrupted (Leclerc et al., 2000, 2003; Moreau and Leclerc, unpublished data). For example, when intact embryos were treated with DHP-sensitive Ca2+ channel blockers, they lacked anterior brain structures (Moreau et al., 1994; Leclerc et al., 1997, 2001). This latter phenotype is similar to the one obtained when the BCNE is surgically removed (Kuroda et al., 2004).

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Are There Ca2+-Sensitive Target Genes?

It has been previously shown (using animal caps) that Ca2+ controls the expression of the immediate early gene c-fos and of two other transcription factors: XlPou2 and Zic3 (Leclerc et al., 1999). While Fos is a ubiquitous transcription factor, XlPou2 and Zic3 are specific to neural determination and primary neural regulators (Witta et al., 1995; Nakata et al., 1997). Leclerc et al. (2000) demonstrated that specific antagonists of DHP-sensitive Ca2+ channels blocked the expression of XlPou2 in response to noggin in animal caps, and dramatically reduced the expression of Zic3 in the whole embryo. Furthermore, in planar explants, the accumulated pattern of Ca2+ correlated with the expression of Zic3, and treatment with the DHP-sensitive Ca2+ channel antagonist, nifedipine, blocked the Ca2+ transients and reduced the level of Zic3 expression (Leclerc et al., 2003). These results suggest that the function of the localized increase in [Ca2+]i that occurs in the dorsal ectoderm during neural induction might be to locally activate genes with proneural activity.

Calcium signaling and cell fate determination during neural induction in amphibian embryos

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5.1 Identification of New Ca2+ Target Genes Involved in Neural Induction To identify new Ca2+ target genes involved in neural induction, a subtractive cDNA library was constructed between untreated (i.e., ectodermal) and short-duration (i.e., 15–45 min) caffeine-treated (i.e., neuralized) animal caps (Batut et al., 2003). Caffeine triggers neural induction via an increase in [Ca2+]i (Moreau et al., 1994), and thus allows the differential isolation of the earliest Ca2+-dependent genes involved in neural determination (Batut et al., 2003). One gene, xPRMT1b, was selected from the 30 early genes identified that were found to be controlled by Ca2+ and expressed in the presumptive neural territories. xPRMT1b is the Xenopus homologue of the mammalian arginine methyltransferase PRMT1 gene (Batut et al., 2005). It was shown that in animal caps, xPRMT1b expression is an early response to an increase in Ca2+ and does not require de novo protein synthesis. Its expression is triggered following the application of noggin or by the inhibition of BMP signaling with tBR (a nonfunctional form of the BMP4 receptor). In addition, these effects are specifically blocked by the Ca2+ chelator, BAPTA. In intact embryos, xPRMT1b is expressed in the neural territories. The early expression of xPRMT1b at the gastrula stage also occurs via a Ca2+-dependent mechanism mediated by the activation of DHP-sensitive Ca2+ channels. Overexpression of xPRMT1b in the neural territories was shown to activate the expression of the neural precursor gene Zic3. In addition, the utilization of a Morpholino-based approach (with an oligonucleotide against xPRMT1b) blocked the expression of the neural markers induced by an increase in Ca2+ such as Zic3 in animal caps, and it impaired anterior neural development in the whole embryo (Batut et al., 2005). Identical phenotypes were obtained with antagonists of DHP-sensitive Ca2+ channels (Leclerc et al., 2000), or by deletion of the BCNE (Kuroda et al., 2004). These results suggest that xPMT1b may provide a direct link between the [Ca2+]i increase and downstream events involved in neural induction.

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Conclusion

Adoption of a neural fate has been, until recently, considered as a permissive event, only requiring the inhibition of BMP signaling. Although this ‘‘by default’’ model has allowed us to understand part of the process of early neurogenesis and epidermal determination at the molecular level, a number of important questions still remain to be addressed. The ‘‘by default’’ model is, however, in conflict with specific data from chick and ascidian embryos, which indicate that neural induction is initiated by FGF signaling in a partly BMP-independent manner (Bertrand et al., 2003; Stern, 2005). The ‘‘by default’’ model also cannot fully explain the inhibition of neuralization triggered by noggin on isolated ectoderm that expresses truncated forms of FGF receptors (Launay et al., 1996). In addition, in intact Xenopus embryos, it has been recently shown that BMP inhibition is required but is not sufficient to trigger neural induction, and that pregastrula FGF signaling is required in the ectoderm for the emergence of neural fates (Delaune et al., 2005). It has, however, been shown that an increase in [Ca2+]i is a necessary and sufficient event to neuralize the ectoderm and that the Ca2+-sensitive gene, xPRMT1b, is expressed in the neural territories in both animal caps and intact embryos, and controls the expression of the early proneural genes. These results thus suggest that Ca2+ plays an instructive rather than permissive role in neural induction. The identification and functional characterization of new Ca2+ target genes, such as xPRMT1b, will help us to further establish the link between Ca2+ influx and neural determination. Several important and as yet unsolved questions have, however, been raised by these new Ca2+-related data; for example, the mechanism by which the DHP-sensitive Ca2+ channels are activated during gastrulation in the dorsal ectoderm is still unknown, as is how noggin can stimulate an influx of Ca2+ through DHP-sensitive Ca2+ channels. In this respect, it is important for us to further consider the relationship between noggin and the FGF receptor. It has been demonstrated, for example, in chick embryonic neurons and in endothelial cells, that the activation of the FGF receptor stimulates the release of arachidonic acid and its metabolites, which in turn activate a Ca2+ influx, perhaps via transient receptor potential channels (TRPs) (Distasi et al., 1995; Antoniotti et al., 2003).

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Calcium signaling and cell fate determination during neural induction in amphibian embryos

Another important question is why the Ca2+ signals are initially generated in the anterior part of the ectoderm? We suggest that this may be explained by the fact that the inducing signal arises via the diffusion of molecules secreted by the dorsal mesoderm. Furthermore, as suggested previously, the anterior region of the blastula stage embryo generating the Ca2+ transients might correspond to the BCNE. The presence of neural inducers, such as noggin, in the blastula ectodermal precursor cells (Kuroda et al., 2004) and the evidence that noggin activates DHP-sensitive Ca2+ channels on animal caps (Leclerc et al., 1999; Batut et al., 2005) also support this hypothesis.

6.1 How Gene Expression During Neural Induction Might Be Controlled by Ca2+ Control of gene expression by Ca2+ very often involves changes in the transactivating properties of transcription factors following the activation of Ca2+-dependent kinases and phosphatases (Dolmetsch et al., 2001; West et al., 2001; Kornhauser et al., 2002; Spotts et al., 2002). However, direct effectors of Ca2+-induced gene expression have also been suggested to exist in the nucleus (Mellstrom and Naranjo, 2001). An important cis-regulatory element, downstream regulatory element (DRE), has been implicated in what can be called excitation- or stimulation-transcription coupling mechanisms (Carrion et al., 1999). The DRE sequence, which is located downstream from the TATA box, is the target of the DRE antagonist modulator (DREAM), an EF-hand Ca2+-binding protein of the recoverin subfamily, which in the absence of Ca2+ binds to the DRE site and represses transcription (Carrion et al., 1999). While, to date, DREAM is the only Ca2+ sensor, which is known to bind specifically to DNA and directly regulate transcription in a Ca2+-dependent manner, there is some evidence suggesting that the promoters of some other neural specific genes might contain putative Ca2+-dependent cis-regulatory elements (I. Ne´ant, unpublished data). Recent findings, including our own, however, suggest an alternative mechanism by which Ca2+ might directly control gene expression. In our new model, the DHP-sensitive Ca2+ channel plays a crucial role in triggering neural induction and neural gene expression. The mechanism, however, that links the activity of this Ca2+ channel to the nucleus is not well understood. The DHP channel may be related to an L-type Ca2+ channel. Recently, it was shown that the C-terminal fragment of an L-type Ca2+ channel translocates to the nucleus, where it binds to a nuclear protein, then associates with an endogenous promoter, and thus regulates transcription of a wide variety of endogenous genes important for neuronal signaling (GomezOspina et al., 2006). This work reports a possible pathway that we suggest might also operate during neural induction, where the activity of DHP-sensitive Ca2+ channels might control the expression of neural genes. Another possible way that Ca2+ might control neural gene expression is through the inhibition of BMP signaling, by acting downstream of Smad phosphorylation. The spatial distribution of activated Smad1 (i.e., phosphorylated Smad1) has been reported to change at the onset of gastrulation (Faure et al., 2000). Prior to gastrulation, phosphorylated Smad1 (which reflects the activation of the BMP4 signaling pathway) is equally distributed on the dorsal and ventral sides of the embryo. In contrast, at late blastula, Smad1 phosphorylation is enriched on the ventral side, and by early gastrulation most of the activated Smad1 is localized to the ventral side (Faure et al., 2000). This correlates with the pattern of [Ca2+]i increase, which starts in the dorsal ectoderm at the blastula stage and is at a maximum during midgastrulation (Leclerc et al., 2000). We suggest that the dephosphorylation of Smad1 in the dorsal ectoderm during gastrulation might be controlled by calcineurin, a Ca2+-/calmodulin-dependent phosphatase 2B. Indeed, in Xenopus embryos, calcineurin is expressed throughout early development (Saneyoshi et al., 2000). Furthermore, the injection of constitutively active mouse calcineurin into a ventral position in Xenopus embryos has been reported to produce a double axis (Nishinakamura et al., 1997). In conclusion, we propose a new model for neural induction in Xenopus embryos to challenge the concept of a simple ‘‘by default’’ mechanism. Our new model integrates the activation of an inductive Ca2+-dependent signaling pathway due to an influx of Ca2+ through DHP-sensitive Ca2+ channels. While our new model proposes that Ca2+ is required for the activation of neural specific genes, it also suggests that epidermal determination occurs when the Ca2+-dependent signaling pathway is inactive. Our new model is schematically illustrated in > Figure 1-4.

Calcium signaling and cell fate determination during neural induction in amphibian embryos

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. Figure 1-4 An alternative model for neural induction in Xenopus laevis embryos. (a) In the absence of neural inducers such as noggin, the BMP signaling pathway leads to the formation of a phosphorylated Smad complex. This complex is translocated into the nucleus where it activates DNA binding proteins that in turn activate the expression of epidermal genes and repress the transcription of neural genes. Furthermore, DHP-Ca2+ channels are not activated and thus xPRMT1b is not stimulated. (b) Neural inducers such as noggin prevent the binding of BMP to its receptors, thus leading to an inhibition of the BMP signal transduction pathway. Evidence indicates that noggin can also directly activate DHP-sensitive Ca2+ channels (Leclerc et al., 1999; Batut et al., 1995). In our updated model, Ca2+ plays a central role, by directly activating Ca2+ target genes, such as xPRMT1b, which in turn control neural gene transcription either directly or via the activation of a Ca2+/calmodulin kinase type II. The Ca2+ signals may also inhibit the BMP-signaling pathway by activating calcineurin, which prevents the phosphorylation of Smads. Activation of DHP-Ca2+ channels may also be achieved by an as-yet unidentified mechanism (indicated by the “?”) resulting from the inhibition of the binding of BMP to its receptor brought about by the presence of noggin. In this model, neural induction is, therefore, not a permissive process but, like epidermal induction, and instructive mechanism

Acknowledgments This work was supported by Centre National de la Recherche Scientifique (CNRS); a joint PICS grant funded by the CNRS; the PROCORE France/Hong Kong Joint Research Scheme (F-HK23/06T) sponsored by the Research Grants Council (RGC) of Hong Kong and the Consulate General of France in Hong Kong; Association pour la Recherche sur le Cancer (ARC); and the following Hong Kong RGC grants: HKUST6214/02M, HKUST6279/03M, HKUST6241/04M, and HKUST6416/06M. This chapter was prepared while A.L.M. was the recipient of a Croucher Foundation Senior Research Fellowship. We also thank Dr Osamu Shimomura for his generous supply of aequorins over the years.

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Calcium signaling and cell fate determination during neural induction in amphibian embryos

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Gualandris L, Rouge P, Duprat AM. 1983. Membrane changes in neural target cells studied with fluorescent lectin probes. J Embryol Exp Morphol 77: 183-200. Gualandris L, Rouge P, Duprat AM. 1985. Target cell surface glycoconjugates and neural induction in an amphibian. J Embryol Exp Morphol 86: 39-51. Keller R, Danilchik M. 1988. Regional expression, pattern and timing of convergence and extension during gastrulation of Xenopus laevis. Development 103: 193-209. Kornhauser JM, Cowan CW, Shaywitz AJ, Dolmetsch RE, Griffith EC, et al. 2002. CREB transcriptional activity in neurons is regulated by multiple, calcium-specific phosphorylation events. Neuron 34: 221-233. Kuroda H, Wessely O, De Robertis EM. 2004. Neural induction in Xenopus: Requirement for ectodermal and endomesodermal signals via Chordin, Noggin, beta-Catenin, and Cerberus. PLoS Biol 2: E92. Launay C, Fromentoux V, Shi DL, Boucaut JC. 1996. A truncated FGF receptor blocks neural induction by endogenous Xenopus inducers. Development 122: 869-880. Leclerc C, Daguzan C, Nicolas MT, Chabret C, Duprat AM, et al. 1997. L-type calcium channel activation controls the in vivo transduction of the neuralizing signal in the amphibian embryos. Mech Dev 64: 105-110. Leclerc C, Duprat AM, Moreau M. 1999. Noggin upregulates Fos expression by a calcium-mediated pathway in amphibian embryos. Dev Growth Differ 41: 227-238. Leclerc C, Lee M, Webb SE, Moreau M, Miller AL. 2003. Calcium transients triggered by planar signals induce the expression of ZIC3 gene during neural induction in Xenopus. Dev Biol 261: 381-390. Leclerc C, Moreau M, Gualandris-Parisot L, Drean G, Canaux S, et al. 1995. An elevation of internal calcium occurring via L-type channels mediate neural induction in the amphibian embryo. Organisation of the Early Vertebrate. Zagris N, Duprat AM, Durston J, editors. New York: Plenum Press; pp 209-226. Leclerc C, Rizzo C, Daguzan C, Neant I, Batut J, et al. 2001. Neural determination in Xenopus laevis. embryos: Control of early neural gene expression by calcium. J Soc Biol 195: 327-337. Leclerc C, Webb SE, Daguzan C, Moreau M, Miller AL. 2000. Imaging patterns of calcium transients during neural induction in Xenopus laevis. embryos. J Cell Sci 113 Pt 19: 3519-3529. Mellstrom B, Naranjo JR. 2001. Ca2+-dependent transcriptional repression and derepression: DREAM, a direct effector. Semin Cell Dev Biol 12: 59-63. Moreau M, Leclerc C, Gualandris-Parisot L, Duprat AM. 1994. Increased internal Ca2+ mediates neural induction

Calcium signaling and cell fate determination during neural induction in amphibian embryos in the amphibian embryo. Proc Natl Acad Sci USA 91: 12639-12643. Nakata K, Nagai T, Aruga J, Mikoshiba K. 1997. Xenopus Zic3, a primary regulator both in neural and neural crest development. Proc Natl Acad Sci USA 94: 1198011985. Nishinakamura R, Matsumoto Y, Uochi T, Asashima M, Yokota T. 1997. Xenopus FK 506-binding protein homolog induces a secondary axis in frog embryos, which is inhibited by coexisting BMP 4 signaling. Biochem Biophys Res Commun 239: 585-591. Okabayashi K, Asashima M. 2003. Tissue generation from amphibian animal caps. Curr Opin Genet Dev 13: 502-507. Saint-Jeannet JP, Foulquier F, Goridis C, Duprat AM. 1989. Expression of N-CAM precedes neural induction in Pleurodeles waltl. (urodele, amphibian). Development 106: 675-683. Saint-Jeannet JP, Huang S, Duprat AM. 1990. Modulation of neural commitment by changes in target cell contacts in Pleurodeles waltl. Dev Biol 141: 93-103. Saint-Jeannet JP, Pituello F, Huang S, Foulquier F, Duprat AM. 1993. Experimentally provoked neural induction results in an incomplete expression of neuronal traits. Exp Cell Res 207: 383-387.

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Saneyoshi T, Kume S, Natsume T, Mikoshiba K. 2000. Molecular cloning and expression profile of Xenopus calcineurin. A subunit(1). Biochim Biophys Acta 1499: 164-170. Sasai Y, De Robertis EM. 1997. Ectodermal patterning in vertebrate embryos. Dev Biol 182: 5-20. Spotts JM, Dolmetsch RE, Greenberg ME. 2002. Time-lapse imaging of a dynamic phosphorylation-dependent protein–protein interaction in mammalian cells. Proc Natl Acad Sci USA 99: 15142-15147. Stern CD. 2005. Neural induction: Old problem, new findings, yet more questions. Development 132: 2007-2021. Takata K, Yamamoto K, Ozawa R. 1981. Use of lectins as probes for analyzing embryonic induction. Roux’s Arch Dev Biol 190: 92-96. Webb SE, Lee KW, Karplus E, Miller AL. 1997. Localized calcium transients accompany furrow positioning, propagation, and deepening during the early cleavage period of zebrafish embryos. Dev Biol 192: 78-92. West AE, Chen WG, Dalva MB, Dolmetsch RE, Kornhauser JM, et al. 2001. Calcium regulation of neuronal gene expression. Proc Natl Acad Sci USA 98: 11024-11031. Witta SE, Agarwal VR, Sato SM. 1995. XIPOU 2, a noggininducible gene, has direct neuralizing activity. Development 121: 721-730.

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Development of the Cerebellum

M. Hashimoto

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16

2 Development of the Cerebellum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Processes of the Cerebellar Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Regionalization During Early Development: A Set Location of the Cerebellum on the Neural Tube . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 The Regional Expression of Transcription Factors Shapes the Cerebellum . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Mediolateral Compartmentalization of the Cerebellum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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3 The Birthdates of PCs Determine the Compartmentalization of the Cerebellum . . . . . . . . . . . . . . . 20 3.1 Adenovirus-mediated Gene Delivery to Birthdate-Specific Populations of Cortical Neurons . . . . 20 3.2 Correlation Between the Birthdate of Purkinje Cells and the M-L Compartmentalization . . . . . . . 21 4

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2009 Springer ScienceþBusiness Media, LLC.

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Development of the cerebellum

List of Abbreviations: Egl, external granular layer; bgal, b-galactosidase

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Introduction

The neuroanatomic topography of the developing cerebellum reveals the formation of distinct structural patterns along its dorsoventral and mediolateral axes. Across the mediolateral axis, parasagittal bands compartmentalize the cerebellar structure into organized stripes of Purkinje cells that are closely associated with their underlying subdivisions of connectivity and function. However, how these parasagittal stripes originate and then organize coherently remains unclear. This review begins with a general outline of cerebellar development and then proceeds to discuss new findings in the origins of cerebellar compartmentalization. Namely, a pulse gene transfer approach employing adenoviral vectors has demonstrated that cerebellar compartmentalization already appears to be determined at the birthdate of Purkinje cells.

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Development of the Cerebellum

2.1 Processes of the Cerebellar Development The complex formation of the brain begins from the neural tube of the embryo. A portion of this neural tube is specialized during development, and then this specific region begins to secrete signaling elements along the antero-posterior and dorso-ventral axes of the neural tube. The secretory elements thus produce positional information along the neural tube. From this nascent positional information, the neural tube establishes the antero-posterior and dorso-ventral axes and the neurodevelopmental regions which follow. For instance, the hindbrain is compartmentalized into rhombomeres, and the telencephalon and diencephalon are also compartmentalized into prosomeres. These early compartments will ultimately develop into the complex brain. The cerebellum also goes through the specific process of regionalization during development (Zervas et al., 2005). The regionalization during the cerebellar development can be divided into the following six steps: 1. The isthmus forms from the boundary between the midbrain and the hindbrain, and it produces signaling factors along antero-posterior axis. These signals induce the cerebellar primordium from rhombomere 1. 2. The consequent expression and activation of several transcription factors begins to regionalize the cerebellar primordium, and this regionalization organizes along the dorso-ventral axis. 3. Two germinal zones, the cerebellar ventricular zone (VZ) and the external granular layer (egl), form in the cerebellum. The cerebellar VZ is located at the roof of the fourth ventricle and mainly produces Purkinje cells (PCs). In contrast, the egl is located at the surface of cerebellum and mainly produces granule cells (GCs). 4. PCs that generated from the cerebellar VZ form striped compartments in the cerebellum by embryonic day (E) 18.5. 5. The GCs generated from the egl migrate radially into the inside of the cerebellum and form the inner granular layer after birth. 6. The afferents, the climbing fibers, and the mossy fibers form specific striped regions in the cerebellum until adulthood. Therefore, a mature neuronal network of the cerebellum is established.

2.2 Regionalization During Early Development: A Set Location of the Cerebellum on the Neural Tube Mouse genetic manipulations are currently revealing the molecular mechanisms of cerebellar morphogenesis (Zervas et al., 2005). Several key molecules are expressed in restricted regions of the embryonic mouse brain and regulate cerebellar development. First, two homeobox genes, Otx2 and Gbx2, are independently expressed in the neural plate early in development. The expression of Otx2 and Gbx2 is observed on the

Development of the cerebellum

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anterior and posterior regions of the neural plate, respectively. The Otx2-positive region is the future forebrain and midbrain, and the Gbx2-positive region is the future hindbrain and spinal cord. The boundaries of Otx2 and Gbx2 gene expression become clearer by E8.5 in mice. Subsequently, the genes of secretory signaling factors Gbx2 and Fgf8 are induced on the Otx2-positive and the Gbx2-positive regions, respectively. In mouse, the patterns of Wnt1 and Fgf8 gene expression tighten at E9.5 when the neural plate closes and forms the neural tube. Wnt1 gene expression localizes discretely to the anterior half of the midbrain–hindbrain boundary. In contrast, Fgf8 gene expression localizes to the posterior half of the midbrain–hindbrain boundary (> Figure 2‐1). Otx2-driven Wnt1 gene expression and Gbx2-driven Fgf8

. Figure 2‐1 Formation of the isthmus (midbrain–hindbrain boundary). The regional expression of transcription factor genes Otx1 (light blue), Wnt1 (red), Fgf8 (yellow), and Gbx2 (green) from the E9.5 mouse embryo is shown. pro, prosencephalon; di, diencephalon; mes, mesencephalon; is, isthmus; r1, rhombomere 1; sc, spinal cord

gene expression provide positional information on the neural tube along the antero-posterior axis and facilitate the regional specification between midbrain and hindbrain. A notable transformation on the midbrain–hindbrain boundary appears at this time. The boundary curves toward the inside of the brain and forms a neck identified as the isthmus. The location of this isthmus corresponds with Fgf8-positive region. As mentioned earlier, the midbrain–hindbrain boundary (the isthmus) arises from the signaling between Otx2/Wnt1 and Gbx2/Fgf8 gene expression, and the secretory factors from the isthmus induce the cerebellum from rhombomere 1.

2.3 The Regional Expression of Transcription Factors Shapes the Cerebellum After E10.5, the isthmic region narrows, the cerebellar primordium expands laterally, and the cerebellar shape becomes clearly distinguishable (> Figure 2‐2). Additionally, the hindbrain that is contiguous to the

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Development of the cerebellum

. Figure 2‐2 Regionalization within the E11.5 cerebellum is formed by the localized expression of functional molecules (a) A schematic view of the posterolateral side of mouse embryo at E11.5. The cerebellar rhombic lip (crl), the hindbrain rhombic lip (hrl), and the roof plate (rp) are colored with green, orange, and yellow, respectively. (b) A schematic view of a sagittal section of the cerebellum (cb) along the red plane shown in A. The ventricular zone (VZ) of the cerebellum, the crl, and the rp are also colored with light blue, green, and yellow, respectively. The transcription factor Lmx1a is locally expressed on the rp, and the TGFb family proteins, Gdf7, BMP6, and BMP7 are secreted from the rp. The cerebellum is compartmentalized into the Math1-positive rp (green region) and the Ptf1a-positive VZ of the cerebellum (light blue region). 3v, third ventricle; 4v, fourth ventricle; egl, external granular cell layer; hb, hindbrain; mb, midbrain; pon, pons

cerebellum separates into two lateral parts along the dorsal midline, and the roof plate, which occupies the area between the cerebellum and the hindbrain. The formation of the roof plate plays an important role in the regionalization of the cerebellum. The roof plate is thin layer of nonneuronal epithelial cells and the future choroid plexus, and the roof plate expresses the LIM homeobox transcription factor Lmx1a. This Lmx1a-positive roof plate secretes the TGFb family proteins Gdf7, BMP6, and BMP7. These proteins induce the expression of the transcription factor Math1 in the cerebellar rhombic lip, a restricted region adjacent to the roof plate in the cerebellum (Alder et al., 1999). The cerebellar VZ adjacent to the Math1-positive cerebellar rhombic lip begins to express the transcription factor Ptf1a (Hoshino et al., 2005). Thus, the cerebellar VZ can be regionalized into a Math1-positive cerebellar rhombic lip and a Ptf1a-positive cerebellar VZ (> Figure 2‐2b, green and blue areas). It is known that Gdf7, BMP6, and BMP7 are secreted from the roof plate and are closely associated with the formation of the Math1-positive cerebellar rhombic lip (Alder et al., 1999; Chizhikov et al., 2006). However, it is not clear what induces the Ptf1a-positive cerebellar VZ because it is evident that roof plate-derived Gdf7, BMP6, and BMP7 are not associated with Ptf1a gene expression (Chizhikov et al., 2006). Recent studies suggest that the neuroepithelial cells from the Math1-positive cerebellar rhombic lip give rise to the glutamatergic neurons including cerebellar GCs and large neurons in the cerebellar nuclei

Development of the cerebellum

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(Machold and Fishell, 2005; Wang et al., 2005). Likewise, the neuroepithelial cells in the Ptf1a-positive cerebellar VZ produce GABAergic neurons including PCs, Golgi cells, basket cells, stellate cells, and small neurons in cerebellar nuclei (Hoshino et al., 2005). Accordingly, as the differentially located expression of Math1 and Ptf1a structurally regionalizes the cerebellum, Math1-positive and Ptf1a-positive regions generate excitatory neurons and inhibitory neurons, respectively. Similarly, the patterns of Math1 and Neurogenin1 gene expression also regionalize the hindbrain rhombic lip. The Math1-positive region is located on the dorsal edge on the hindbrain rhombic lip, and the Neurogenin1-positive region is contiguous to this Math1-positive region. Math1-positive neuroepithelial cells generate neurons of the pons, the lateral reticular nucleus, and the accessory cuneate nucleus. Neurogenin1-positive neuroepithelial cells generate neurons of the inferior olive (Landsberg et al., 2005). Math1-positive cells generated from the cerebellar rhombic lip migrate anteriorly along the outer surface of the cerebellum, then form the egl. At this stage, two different germinal layers, the external GC layer (Math1-positive cell group) and the cerebellar VZ (Ptf1a- positive cell group), are formed within the cerebellum. The PCs from the cerebellar VZ migrate from the ventricular side to the surface of the cerebellum and then form the PC layer. In contrast, the GCs from the external GC layer move from the external GC layer (> Figure 2‐2b, egl) to the ventricular side and form the inner GC layer.

2.4 Mediolateral Compartmentalization of the Cerebellum The adult cerebellum is functionally compartmentalized along the mediolateral (M-L) axis. Clear regionalization is observed in the neural circuit of cerebellum. Neurons of the inferior olivary nuclei project their axons (climbing fibers) toward cerebellar PCs. Interestingly, the climbing fibers projected from the subset of neurons in a subnucleus of the inferior olive form specific striped compartments in the cerebellum (> Figure 2‐3). The striped regions formed by the climbing fibers show a symmetric distribution. In addition, the subset of PCs belonging to the striped region projects their axons to a specific deep cerebellar nucleus (> Figure 2‐3). As a result, the subnucleus of inferior olivary nuclei – PCs – the deep cerebellar nucleus forms zone-to-zone innervations (Voogd and Glickstein, 1998). Furthermore, physiological studies confirm that the striped regionalization from the climbing fibers codes a somatosensory map concerning the body movement (Voogd and Glickstein, 1998; Apps and Garwicz, 2005). Therefore, the M-L compartments in the cerebellum are thought to be important as the functional units of the cerebellum. Molecular studies indicate that a variety of molecular markers are expressed in specific subsets of M-L compartments. For instance, gene expression of L7/pcp2 (a genetic marker of cerebellar PCs), engrailed (En)1, En2, Pax2, and Wnt7B (mammalian homologues of Drosophila segment polarity genes) appears in specific striped-regions. In addition, the expression of Zebrin II (aldolase C) also shows a specific striped-pattern. Interestingly, En2-deficient mice show an abnormal morphology of cerebellum (Kuemerle et al., 1997) and the striped pattern of Zebrin II closely correlates to the striped region formed by the climbing fibers (Leclerc et al., 1988; Sugihara et al., 2000; Sugihara and Quy, 2007). However, the striped pattern of their expression is not stable. The striped-patterns of L7/pcp2, En1, En2, Pax2, and Wnt7B gene expression initially appear from E15.5, but then the striped-expression disappear shortly after birth. In addition, all PCs express Zebrin II by the first week after birth, but the expression of Zebrin II are gradually changed to a striped-pattern within the cerebellum until 20 days after birth (Leclerc et al., 1988). The striped pattern of Zebrin II is entirely different from the L7/pcp2, En1, En2, Pax2, and Wnt7B gene expression patterns. So, it is difficult to examine the process of the cerebellar regionalization because the forming process with the expression of these marker genes is difficult to trace. The striped compartments in the cerebellum are observed not only at the neural circuit and neurophysiologic level, but also at the cellular and molecular levels. These observations suggest that cerebellar M-L compartments are the basic modular structures from which cerebellar functions are performed. However, the mechanisms that initiate and establish such cerebellar M-L compartmentalization remain unclear.

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Development of the cerebellum

. Figure 2‐3 M-L compartmentalization of the cerebellar neural projections. Neurons from a specific subnucleus of the inferior olive nucleus project their axons (climbing fibers) into the contralateral cerebellar cortex. The climbing fiber terminals organize into striped, compartmentalized regions within the cerebellum along the M-L axis. Additionally, the PCs within each M-L compartment project their axons into a particular region of the cerebellar deep and vestibular nuclei. The neural projections from the inferior olivary nucleus, through the M-L compartments of the cerebellum, and to the cerebellar deep and vestibular nuclei establish zone-to-zone innervations. As such, the M-L compartments revealed by the cerebellar neural projection are thought as the structural unit of the cerebellum

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The Birthdates of PCs Determine the Compartmentalization of the Cerebellum

3.1 Adenovirus-mediated Gene Delivery to Birthdate-Specific Populations of Cortical Neurons During cortical development, early-born neuronal progenitors migrate to the inner cortical layers, and lateborn neuronal progenitors migrate to the outer superficial layers. This timing establishes an inside-out gradient of cortical development, and the progenitor cells differentiate to acquire layer-specific morphologies, connections, and function. However, the mechanisms involved in such neuronal fate remain unclear, especially because it is difficult to identify and to track a particular subset of progenitor cells. Thus, in order to investigate the laminar development of cortical structures in the brain, it is first critical to distinguish a particular subset of progenitor cells that share the same neuronal birthdate from other progenitor cells with different birthdates. Using replication-defective adenoviral vectors, we have successfully performed a ‘‘pulse gene transfer’’ approach to deliver an exogenous gene into restricted subpopulations of neuronal progenitor cells and in a birthdate-specific manner (Hashimoto and Mikoshiba). Briefly, the adenoviral vector AdexCAG-NL-LacZ, a viral vector designed for nucleus-targeted b-galactosidase (bgal) expression, is injected into the midbrain ventricles of mouse embryos between E10.5 and E14.5 via exo utero surgery (> Figure 2‐4). Remarkably, the

Development of the cerebellum

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. Figure 2‐4 Construction of the adenoviral vector AdexCAG-NL-LacZ and exo utero surgery for the adenoviral injection. (a) A schematic drawing of the adenoviral vector AdexCAG-NL-LacZ. AdexCAG-NL-LacZ expresses nucleartargeted b-galactosidase (bgal) under the control of the robust and constitutively active CAG promoter. The adenoviral vector is based on human adenovirus type 5 (Ad5) and is replication-incompetent for lack of E1A and E1B that are essential for the adenoviral replication. (b) A schematic drawing of exo utero surgeries. The AdexCAG-NL-LacZ adenoviral vector was injected into the midbrain ventricle (m) of mouse embryos with a glass pipette (p). The procedure of exo utero surgery is described in previous reports (Muneoka et al., 1986; Hashimoto and Mikoshiba, 2003; 2004). 3, third ventricle; 4, fourth ventricle; em, extra membrane; t, telencephalon; scale bar, 1mm

adenoviral vector infects specific cohort subsets of birthdate-related progenitor cells, and these virally infected cohorts proceed to develop normally into cortical neurons and formed canonical cortical layers in the expected inside-out manner (> Figure 2‐5). Similar to the tritiated thymidine and bromodeoxyuridine (BrdU) pulse-labeling approach for neuronal birthdate analyses, this adenovirus approach enables the examination of distinct subsets of progenitor cells that share the same neuronal birthdate (Hashimoto and Mikoshiba, 2004). In addition, this gene transfer approach will also provide the future means to edit the molecular background of differentiating progenitors in order to address the mechanisms that may determine neuronal fate within each cortical layer. This adenovirus-mediated gene transfer technique will illuminate the molecular properties of birthdaterelated progenitor cell subpopulations and the mechanisms by which the laminar structures of the cortex are formed in the brain.

3.2 Correlation Between the Birthdate of Purkinje Cells and the M-L Compartmentalization The advantage of this timed adenovirus gene transfer approach can be applied to the study of the cerebellar development and regionalization (Hashimoto and Mikoshiba). When AdexCAG-NL-LacZ is injected into the midbrain ventricle of the mouse embryo, it also infects the progenitor cells on the surface of the fourth ventricle and effectively transfers the nuclear-targeted bgal into the progenitor cells. Interestingly, the injection of AdexCAG-NL-LacZ into embryos at E10.5, E11.5, and E12.5 reveals that successfully infected bgal-positive progenitor cells develop normally and differentiate into cerebellar PCs. The result is consistent with the neuronal birthdate analysis of PCs (Altman and Bayer, 1997) and a report indicating that cerebellar PCs are generated from Ptf1a-positive progenitor cells (> Figure 2‐2, blue area) on the surface of the fourth ventricle (Hoshino et al., 2005). By using adenoviral vectors, we can efficiently transfer a foreign gene into the progenitor cells of PCs in a neuronal birthdate-specific manner and observe the native behavior of each

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Development of the cerebellum

. Figure 2‐5 Neuronal birthdate-specific gene transfer via adenoviral vector. Abbreviations in the format ‘‘E12.5:P20’’ are used throughout this text. The left side (‘‘E12.5’’) indicates the embryonic stage at which the adenoviral vector was injected, and the right side (‘‘P20’’) is the age at which the mice were analyzed. The bgal-positive cells were clearly localized in layers V, IV, and II/III of the E12.5:P20, E13.5:P20, and E14.5:P20 neocortices, respectively. These results indicate that early-infected progenitor cells (e.g., E12.5) came to lie in the inner layer (e.g., layer V), while late-infected progenitor cells (e.g., E14.5) migrated to the outer layer (e.g., layer II/III). Thus, the virally infected progenitor cells revealed an ‘‘inside-out’’ developmental gradient. The neuronal birthdate-specific gene transfer mediated by the adenoviral vector can be observed in other regions in the brain as well. The results are parallel to pulse-labeling with tritiated thymidine and BrdU. IV–VI, the layer number in the cerebral cortex; A, dorsal view; B, ventral view; C, lateral view; H, hippocampus; Apm, posteromedial cortical amygdaloid nucleus; Apl, posterolateral cortical amygdaloid nucleus; HT, hypothalamus; PIR, piriform cortex

cohort of PCs that share the same neuronal birthdate. Surprisingly, the PCs that shared the same birthdate form specific subsets of M-L compartments in the cerebellum (> Figures 2‐6 and > 2‐7). When AdexCAG-NL-LacZ is injected into the E11.5 embryo, the cerebellum at P20 (E11.5:P20; arrowheads in > Figure 2‐6a) indicates five striped regions that are negative for bgal (Hashimoto and Mikoshiba, 2003). In contrast, if AdexCAG-NL-LacZ is injected into the E12.5 embryo, the P 20 cerebellum (E12.5:P20; > Figure 2‐6b) indicates three striped regions that are positive for bgal. The patterns of the striped regions are entirely different between E11.5:P20 (> Figure 2‐6a) and E12.5:P20 (> Figure 2‐6b) cerebella. Each subset of M-L compartments displays nested and, in part, mutually complementary patterns, and these patterns is unchanged from the late embryonic stage to adulthood. The M-L compartments did not change throughout the life of the mouse (Hashimoto, unpublished data). In addition, the bgal-positive M-L compartments formed by E12.5-born PCs (> Figure 2‐6b) are similar to the pattern of the striped regions established by climbing fibers (Sugihara et al., 2000).

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. Figure 2‐6 Purkinje cells sharing the same neuronal birthdate form a specific subset of M-L compartments in the cerebellum. AdexCAG-NL-LacZ was injected into the midbrain ventricle of mouse embryos on E11.5 and E12.5. Twenty days after birth (P20), each manipulated brain was stained by whole-mount for bgal. High bgal activity was observed in PCs in the E11.5:P20 (a) and E12.5:P20 (b) cerebella. In contrast, there was little bgal activity in the E13.5:P20 and E14.5:P20 cerebella (> Figure 2‐5). bgal-positive PCs in the E11.5:P20 and E12.5:P20 cerebella formed M-L compartments. The M-L compartments continued to develop without interruption from the posterior into the anterior lobe. The pattern of bgal-positive M-L compartments in the E11.5:P20 cerebellum was completely different from that in the E12.5:P20 cerebellum. (a) The arrowheads indicate five bgal-negative bands. Interestingly, PCs in the anterior half of the paraflocculus (PF) are positive for bgal. (b) Three bgal positive bands are observed

The fundamental patterns of the M-L compartments formed by birthdate-related PCs are established at E18.5 (Hashimoto and Mikoshiba, 2003). To clarify the correlation between PC birthdates and the cerebellar M-L compartmentalization, the M-L compartments formed by E10.5-, E11.5-, and E12.5-born PCs are compared at E18.5 cerebella (E10.5:E18.5, E11.5:E18.5, and E12.5:E18.5 in > Figure 2‐7, respectively). The E10.5:E18.5 cerebellum shows compartments 3, 6, and 8, but compartment 8 is ambiguous because it is positive for bgal in the near lateral view, but not in the dorsal view (‘‘E10.5:E18.5’’ line in > Figure 2‐7). In the E11.5:E18.5 cerebellum, compartments 2, 4, 5, and 8 are clearly demarcated (‘‘E11.5:E18.5’’ line in > Figure 2‐7), and in the E12.5:E18.5 cerebellum, compartments 1, 2, 5, and 7 can be seen clearly (‘‘E12.5:E18.5’’ line in > Figure 2‐7). Compartments 3 and 6 in the E10.5:E18.5 cerebellum compensate for gaps of M-L compartments in the E11.5:E18.5 and E12.5:E18.5 cerebella. Our results indicate that the PC progenitor cells are already predestined to form specific subsets of M-L compartments on their neuronal birthdates between E10.5 and E12.5. This reviewed study (Hashimoto and Mikoshiba,

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Development of the cerebellum

. Figure 2‐7 Comparison of specific M-L compartment patterns formed by birthdate-related Purkinje cells on E18.5. The posterior (p), dorsal (d) and near lateral (l) views of E10.5:E18.5, E11.5:E18.5 and E12.5:E18.5 cerebella are indicated in each line. Each cerebellum showed a specific pattern of M-L compartments. The M-L compartment positions are defined and named from 1 to 8, as indicated at the top of each line. The M-L compartment boundaries are indicated by dotted lines. The spatial relationship of the M-L compartments in E10.5:E18.5, E11.5:E18.5, E12.5:E18.5 cerebella are summarized in the bottom line. Scale bar, 1 mm

2003) represents the first such direct observation of PC development. The birthdates of PCs determine the compartmentalization of the cerebellum.

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Conclusion

I have confirmed that the neuronal birthdate of PCs closely regulates the M-L compartmentalization of the cerebellum. I hypothesize that the pattern of cerebellar compartmentalization is generated through two

Development of the cerebellum

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independent steps. The first step is a ‘‘generation step’’ in the VZ, and the second step is a ‘‘subdividing step’’ in the mantle zone (MZ). In the initial generation step, the first PCs to be generated on E10.5 immediately acquire their cell fate, as characterized by the expression of specific molecules. On the next day (E11.5), newly generated E11.5 PCs are exposed to extrinsic signals from the E10.5 PCs. This signaling consequently affects the cell fate of the newly generated E11.5 PCs such that they acquire a distinct cellular and molecular identity that distinguishes them from the E10.5 PCs. In the subsequent subdividing step, each subset of PCs leaves the VZ and migrates into the MZ without mingling with each other. These subsets are exposed to extrinsic cues such as cell–cell recognition molecules that play a key role in subdividing them into distinct M-L compartments. The PCs choose migration paths or regions according to their identities and to their extrinsic signals. Consequently, PCs are subdivided into discrete groups, and the nested M-L compartments are formed. However, the molecular mechanisms regulating the formation of the M-L compartments remain unclear. The adenoviral vector approach will help to understand the molecular mechanisms involved in cerebellar M-L compartmentalization and in the physiological functions of each M-L compartment formed by the Purkinje cells that share the same neuronal birthdate.

References Alder J, Lee KJ, Jessell TM, Hatten ME. 1999. Generation of cerebellar granule neurons in vivo by transplantation of BMPtreated neural progenitor cells. Nat Neurosci 2: 535-540. Altman J, Bayer SA. 1997. Development of the Cerebellar System: In Relation to Its Evolution, Structure, and Functions. Boca Raton: CRC Press. Apps R, Garwicz M. 2005. Anatomical and physiological foundations of cerebellar information processing. Nat Rev Neurosci 6: 297-311. Chizhikov VV, Lindgren AG, Currle DS, Rose MF, Monuki ES, et al. 2006. The roof plate regulates cerebellar celltype specification and proliferation. Development 133: 2793-2804. Hashimoto M, Mikoshiba K. 2003. Mediolateral compartmentalization of the cerebellum is determined on the ‘‘birth date’’ of Purkinje cells. J Neurosci 23: 11342-11351. Hashimoto M, Mikoshiba K. 2004. Neuronal birthdate-specific gene transfer with adenoviral vectors. J Neurosci 24: 286-296. Hoshino M, Nakamura S, Mori K, Kawauchi T, Terao M, et al. 2005. Ptf1a a bHLH transcriptional gene defines GABAergic neuronal fates in cerebellum. Neuron 47: 201-213. Kuemerle B, Zanjani H, Joyner A, Herrup K. 1997. Pattern deformities and cell loss in Engrailed-2 mutant mice suggest two separate patterning events during cerebellar development. J Neurosci 17: 7881-7889. Landsberg RL, Awatramani RB, Hunter NL, Farago AF, DiPietrantonio HJ, et al. 2005. Hindbrain rhombic lip is

comprised of discrete progenitor cell populations allocated by Pax6. Neuron 48: 933-947. Leclerc N, Gravel C, Hawkes R. 1988. Development of parasagittal zonation in the rat cerebellar cortex: MabQ113 antigenic bands are created postnatally by the suppression of antigen expression in a subset of Purkinje cells. J Comp Neurol 273: 399-420. Machold R, Fishell G. 2005. Math1 is expressed in temporally discrete pools of cerebellar rhombic-lip neural progenitors. Neuron 48: 17-24. Muneoka K, Wanek N, Bryant SV. 1986. Mouse embryos develop normally exo utero. J Exp Zool 239: 289-293. Sugihara I, Bailly Y, Mariani J. 2000. Olivocerebellar climbing fibers in the granuloprival cerebellum: Morphological study of individual axonal projections in the X-irradiated rat. J Neurosci 20: 3745-3760. Sugihara I, Quy PN. 2007. Identification of aldolase C compartments in the mouse cerebellar cortex by olivocerebellar labeling. J Comp Neurol 500: 1076-1092. Voogd J, Glickstein M. 1998. The anatomy of the cerebellum. Trends Neurosci 21: 370-375. Wang VY, Rose MF, Zoghbi HY. 2005. Math1 expression redefines the rhombic lip derivatives and reveals novel lineages within the brainstem and cerebellum. Neuron 48: 31-43. Zervas M, Blaess S, Joyner AL. 2005. Classical embryological studies and modern genetic analysis of midbrain and cerebellum development. Curr Top Dev Biol 69: 101-138.

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Regulation of Axon Formation

T. Yoshimura . N. Arimura . K. Kaibuchi

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

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Processes of Neuronal Polarization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

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Specification by Intrinsic Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

4 4.1 4.2 4.3 4.4

Neuronal Polarity-Regulating Molecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . PI3-kinase/Akt/GSK-3b Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of CRMP-2 by GSK-3b . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Positive Feedback Loop . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rap1B, H-Ras, and R-Ras . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Conclusion and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35

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Abstract: A mature neuron is typically polarized with a single long axon and several dendrites. After the birth and differentiation of a neuron, a neuron breaks its previous symmetry and establishes an axon and dendrites. Neuronal polarization occurs when one of the multiple immature neurites emerging from the cell body elongates rapidly. This neurite becomes the axon, whereas the remaining immature neurites become dendrites. What are the molecular mechanisms specifying the axon in the initial events? Here we provide an overview of recent progress into the study of axon formation. List of Abbreviations: APC, adenomatous polyposis coli; aPKC, atypical protein kinase C; CRMP-2, collapsin response mediator protein-2; GEF, guanine nucleotide exchange factor; GSK-3b, glycogen synthase kinase-3b; ILK, integrin-linked kinase; NgCAM, neuron-glia cell adhesion molecule; PI3-kinase, phosphatidylinositol 3-kinase; PIP3, phosphatidylinositol 3,4,5-triphosphate; PTEN, phosphatase and tensin homolog deleted on chromosome 10; siRNA, short interfering RNA; Sra-1, specifically Rac1-associated protein 1; STEF, Sif- and Tiam1-like exchange factor; Tiam1, T-lymphoma invasion and metastasis 1

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Introduction

Neuronal polarity is essential for unidirectional signal flow from somata or dendrites to axons. A model system for studying neuronal polarity, in cultured hippocampal neurons, was pioneered by Banker more than 20 years ago (Craig and Banker, 1994). Dotti et al. (1988) precisely observed this differentiation process and divided the morphological events into five stages. Neuronal polarization occurs from stage 2 to stage 3. The first step in neuronal polarization is initial axon formation. At the start of the twenty-first century, studies of axon formation have made advances with the molecular biological approach (Arimura and Kaibuchi, 2007). We are only now beginning to understand the molecular mechanisms involved in the establishment of neuronal polarity. Some pieces of the neuronal polarity puzzle have been solved. There are two major signaling cascades in neuronal polarization: the phosphatidylinositol 3-kinase (PI3-kinase)/Akt (also known as PKB)/glycogen synthase kinase-3b (GSK-3b)/collapsin response mediator protein-2 (CRMP-2) pathway, and the positive feedback loop composed of Rho family small GTPases and the Par3/Par6/atypical protein kinase C (aPKC) complex. Two major signaling cascades downstream of the PI3-kinase play a central role in neuronal polarization.

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Processes of Neuronal Polarization

The ability of cells to polarize is essential for complex biological activities such as the organization of the nervous system. A well-established model for studying neuronal polarity is a hippocampal neuron. Banker and his colleagues developed a culture system of embryonic hippocampal neurons and described in detail the morphological changes that occur during polarization (Dotti et al., 1988). They divided the morphological events into five stages (> Figure 3‐1). First, after being dissociated from embryonic rat brains, hippocampal neurons form lamellipodia and several thin filopodia (stage 1). After several hours, the neurons form several immature neurites, so-called minor processes (stage 2). These neurites are morphologically equal and undergo repeated, random growth and retraction. Half a day after plating, one of these minor processes begins to extend rapidly, becoming much longer than the other neurites (stage 3). This extended process becomes an axon; the other immature neurites continue to undergo brief spurts of growth and retraction, maintaining their net length, for up to a week, when they then become mature dendrites (stage 4). During this process, dendrites become thicker and shorter than axons and begin to establish dendritic components and construct premature dendritic spines (stage 5). When the axon and dendrites are mature, neurons form synaptic contacts that enable the transmission of electrical activity. Although culturing hippocampal neurons has became the most popular tool to monitor neuronal polarity, the morphological changes seen in immature neurons in different brain areas, tissues, and organisms are not necessarily identical in vivo and in vitro. It is possible that the extracellular signals decide the direction of neurons in the early stages. Little is known about extracellular cues that govern

Regulation of axon formation

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. Figure 3‐1 Processes of neuronal polarization in cultured hippocampal neurons. Cultured pyramidal neurons from a rodent hippocampus acquire their characteristic polarized morphology in five well-defined stages (reproduced from Dotti et al. (1988), with permission). During development, neurons acquire their polarity from stage 2 to stage 3. Certain molecular biological manipulations in stage 2 neurons affect axon formation. For example, overexpression of collapsin response mediator protein-2 (CRMP-2) induces multiple axons, whereas depletion of CRMP-2 by short interfering RNA inhibits axon formation

neuronal polarity in vivo. Some studies have shown that extracellular substrate molecules can govern which neurite becomes an axon, depending on the substrate preference of neurite elongation (Esch et al., 1999). When neurons are cultured on substrates patterned with stripes of poly-L-lysine and either laminin or neuron-glia cell adhesion molecule (NgCAM), undifferentiated immature neurites attach on both substrates equally, but axons form preferentially on laminin or NgCAM. These observations suggest that the signals produced by the contact of laminin or NgCAM with adhesion molecules, such as integrins, cause the rapid neurite growth and are sufficient to induce axon formation. This rapid axon formation is also observed when an immature neurite in stage 2 comes in contact with laminin-coated beads (Menager et al., 2004). Recently, it was shown that Wnt5a, a noncanonical Wnt, activated aPKC via Dishevelled and promoted axon specification (Zhang et al., 2007). Thus signaling cascades accelerated by an extracellular matrix may initiate neurite growth and axon formation, and certain extracellular cues may determine axon or dendrite fate during physiological development.

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Specification by Intrinsic Signaling

Because neurons develop polarity in culture without any directional gradients of extracellular cues, neurons appear to have an internal polarization program. How is a single axon specified among equally potential neurites in cultured hippocampal neurons? A possible scenario is that signaling factors (morphogens) form positive and negative feedback loops that can interchangeably affect each other (> Figure 3‐2;

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Regulation of axon formation

. Figure 3-2 A model for axon specification in neuronal polarization. (a) Positive and negative feedback regulation leads to symmetry breaking in a neuron between stage 2 and stage 3. During stage 2, immature neurites extend and retract randomly to maintain their overall length. Arrows represent negative feedback signals that are generated at each growth cone, propagate throughout the cells, and counteract the positive feedback loops in other neurites. When the balance between positive and negative signals is upset (in the transition from stage 2 to stage 3) by extracellular signals, (auto-)activation of receptors or adhesion molecules, and recruitment of signaling molecules, one neurite elongates rapidly. Continuous elongation is supported by a positive feedback loop and sustains the activation cycle. The inhibitory signals that mutually antagonize neurite extension (negative feedback signals) are progressively generated at the growing axon more than at the other neurite (a thicker arrow) and interfere with the specification into axons. (b) The site of axon regeneration after axotomy at a site (1 or 2) is dependent on the relative length of the cut axon compared with the other neurites. If more than 10 mm longer than the other neurites, (1) the axon regenerates again. Otherwise, (2) any neurite can generate a new axon after a long latency

Andersen and Bi, 2000). Neurite extension and retraction are controlled by positive or negative signaling molecules, respectively. The extension of neurites is driven by four main steps: an increase in the amount of plasma membrane by vesicle recruitment and fusion; the local concentration and activation of signaling molecules, such as PI3-kinase and Rho GTPase, and their receptors; an increase in the dynamics of actin filaments; and the enhancement of microtubule formation.

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Immediately after a small amount of growth, the opposite reaction is induced; microtubule catastrophe (collapse), a decrease in the dynamics of actin filaments, and enhanced endocytosis occur. The study of neurite motility indicates that these signaling cascades do indeed form an extension–retraction cycle in the immature neurites (> Figure 3‐2). Moreover, each neurite seems to release negative feedback signals that feed back to the cell body or metabolize molecules that are required for axon specification, either of which might antagonize axon specification in other neurites (Andersen and Bi, 2000). Before polarization, therefore, positive and negative signals seem to be intricately balanced. When this balance is broken by a positive cue, this leads to activation of a continuous self-activation system (positive feedback loop), and a single immature neurite elongates to become an axon (> Figure 3‐2; Goslin and Banker, 1989; Andersen and Bi, 2000). Concurrently, this self-activation system creates strong negative feedback signals that prevent other neurites from forming a second axon (Andersen and Bi, 2000). Direct evidence for the existence of negative feedback signals comes from the following observation: growth/ elongation of a single neurite driven by laminin-coated beads is inhibited by the contact, and subsequent elongation, of a second neurite by laminin beads (Menager et al., 2004). This result indicates the potential existence of negative feedback signals that propagate from the tips of neurites to the cell body; such signals could be induced by the activation of transmembrane receptors. This hypothesis addresses the issue of why only a single neurite is selected to become an axon and indicates that promoting neurite elongation might lead to neuronal polarization. The specification of the axon is thought to depend on its length relative to the other minor processes (Bradke and Dotti, 2000a). Intracellular mechanisms that help to enhance neurite and axon outgrowth evidently require reorganization of cytoskeletons, including actin filaments and microtubules (Arimura and Kaibuchi, 2007). A highly dynamic area is located at the tips of growing axons, where drastic rearrangements of actin filaments and microtubules occur during neurite elongation (Bradke and Dotti, 2000a; Baas and Buster, 2004). Actin instability is higher in one of the unpolarized neuron’s neurite growth cones, and application of the actin-depolymerizing drug cytochalasin D to stage 2 neurons causes multiple axons, implying that reorganization of actin filaments is necessary for axon formation (Bradke and Dotti, 1999). Actin disassembly by cytochalasin B allows microtubules to extend distally into the peripheral region of the growth cones and leads to rapid neurite growth (Forscher and Smith, 1988). Microtubule assembly occurs in the cell body and the growth cone (Brown et al., 1992, 1993). The microtubule array in the neurite or axon is formed through two mechanisms: the transport of microtubule polymer, and microtubule assembly at the plus ends of the microtubules. Both mechanisms appear to contribute to axon outgrowth (Baas, 1997). These findings suggest that actin filaments restrict the protrusion of microtubules. Dynamic actin filaments appear to allow the enhanced polymerization and/or transport of microtubules, resulting in promoted neurite elongation, followed by axon formation (Bradke and Dotti, 2000a). Even after the axon is specified, other neurites have the potential to change their fate during development. Banker and his colleagues reported that transecting the axons of hippocampal neurons early in development could cause the polarity to alternate; an immature neurite starts to become an axon, instead of the transected axon (Dotti and Banker, 1987; Goslin and Banker, 1989). This change depends on the difference in the length of the cut axon compared to the neurites (> Figure 3‐2). If the cut axon is at least 10 mm shorter than the other minor processes, the shortened axon becomes a dendrite, and the longest minor process begins to grow rapidly and eventually becomes an axon. Axon-dendrite plasticity is known to be retained in stage 4 neurons (Bradke and Dotti, 2000b), indicating that to some degree, the identities of axons and dendrites seem to be flexible during development.

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Neuronal Polarity-Regulating Molecules

Many intracellular molecules that compose the signaling networks may regulate axon formation in the early developmental stages. Toriyama et al. (2006) reported that 277 proteins are consistently upregulated during the transition from stage 2 to stage 3 by proteome analyses of cultured hippocampal neurons using highly sensitive large-gel two-dimensional electrophoresis. One of the proteins was analyzed by mass spectrometry

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and named Shootin1 because of a novel protein. Overexpression of Shootin1 induces multiple axons, whereas repressing Shootin1 expression inhibits polarization. These results suggest that Shootin1 regulates axon formation (Toriyama et al., 2006).

4.1 PI3-kinase/Akt/GSK-3b Pathway Recent studies have shown the importance of PI3-kinase and its lipid product [phosphatidylinositol 3,4,5triphosphate (PIP3)] in determining and maintaining internal polarity in neurotrophils and dictyostelium (Iijima et al., 2002). Several groups, including ours, reported that local activation of PI3-kinase and accumulation of PIP3 at the tip of one of the immature neurites are important for axon specification and elongation (> Figures 3‐1 and > 3‐3; Shi et al., 2003; Menager et al., 2004). PI3-kinase inhibitors, such as LY294002, delay the transition of neurons from stage 1 to stage 3, affecting both axon formation and elongation (Shi et al., 2003; Menager et al., 2004). Local contact of immature neurites with an extracellular matrix, such as laminin, induces rapid production of PIP3 at the tip of the neurite through the action of PI3-kinase, and PIP3 is involved in axon specification, possibly by stimulating elongation of an immature neurite (Menager et al., 2004). We previously reported that PIP3 seems to be transported toward the . Figure 3‐3 Signaling cascades in axon formation. In one immature neurite (the future axon), extracellular matrix activates phosphatidylinositol 3-kinase through interaction with adhesion molecules or receptors, thereby producing phosphatidylinositol 3,4,5-triphosphate (PIP3). Accumulated PIP3 drives two major signaling cascades: the Akt/ glycogen synthase kinase-3b/CRMP-2 pathway and the positive feedback loop composed of Cdc42, the Par complex, and Rac1. These signaling cascades regulate cytoskeletons, endocytosis, protein trafficking, and transcriptions to promote neurite elongation and determine axon or dendrite fate

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contact site (Menager et al., 2004). In fact, Horiguchi et al. (2006) recently demonstrated that a complex of guanylate kinase-associated kinesin and a PIP3-interacting protein, PIP3BP, transports PIP3 to the neurite end. The phosphatase and tensin homolog deleted on chromosome 10 (PTEN) is a lipid and protein phosphatase that functions in an opposite manner from PI3-kinase by dephosphorylating PIP3 (Maehama and Dixon, 1998). Overexpression of PTEN inhibits axon formation, whereas knockdown of PTEN by short interfering RNA (siRNA) induces formation of multiple axons (Jiang et al., 2005). PI3-kinase activates Akt by phosphorylation of Akt at Thr-308 and Ser-473 via PIP3, phosphoinositidedependent kinase, and integrin-linked kinase (ILK; Scheid and Woodgett, 2001; Hannigan et al., 2005). Constitutively active myristoylated Akt leads to formation of multiple axons (Jiang et al., 2005). GSK-3b is known to be constitutively active. Activated Akt and ILK phosphorylate GSK-3b at Ser-9 and inactivate its kinase activity (Grimes and Jope, 2001; Hannigan et al., 2005). GSK-3b relays signals from PTEN and Akt, and the decreased activity of GSK-3b is required for neuronal polarization (Jiang et al., 2005; Yoshimura et al., 2005). Inhibition of GSK-3b by GSK-3 inhibitors (LiCl, SB216763, and SB415286) or hairpin siRNA induces formation of multiple axons, whereas overexpression of constitutively active GSK-3b inhibits axon formation (Jiang et al., 2005; Yoshimura et al., 2005). Recently, it was shown that local Akt degradation mediated by the ubiquitin-proteasome system is important in determining neuronal polarity (Yan et al., 2006). Akt is present in both the cell body and multiple immature neurites of stage 2 neurons. Preferential degradation of putative future dendritic Akt is mediated by the ubiquitin-proteasome system from stage 2 to stage 3, and in stage 3 neurons, Akt is localized in the cell body and one axonal tip. There is more inactivated and phosphorylated GSK-3b at Ser-9 in the tips of axons than in the tips of future dendrites in stage 3 neurons (Jiang et al., 2005). At the tips of nascent axons, Akt phosphorylates and inactivates GSK-3b. These results indicate the significance of the PI3-kinase/Akt/GSK-3b pathway in neuronal polarity (> Figures 3‐1 and > 3‐3).

4.2 Regulation of CRMP-2 by GSK-3b CRMP-2 is one of at least five isoforms and is highly and exclusively expressed in the developing nervous system (Goshima et al., 1995; Wang and Strittmatter, 1996; Arimura et al., 2004). Mutations in the unc-33 gene, a Caenorhabditis elegans homolog of CRMPs, lead to severely uncoordinated movement and abnormalities in the guidance of axons of many neurons (Hedgecock et al., 1985). We showed that GSK-3b determines axon or dendrite fate through phosphorylation of CRMP-2 (Yoshimura et al., 2005). CRMP-2 is enriched in the growing axon of hippocampal neurons, and overexpression of CRMP-2 induces multiple axons, whereas inhibition of CRMP-2 functions impairs axon formation (Inagaki et al., 2001). CRMP-2 interacts with tubulin heterodimers and promotes microtubule assembly in vitro (Fukata et al., 2002). CRMP-2 also regulates endocytosis of specific adhesion molecules, including L1, through interaction with Numb (Nishimura et al., 2003) and reorganization of actin filaments acting through specifically Rac1-associated protein 1 (Sra-1; Kawano et al., 2005). Truncated Kinesin-1 selectively accumulates in only (future) axons in the early and late stages (Nakata and Hirokawa, 2003; Jacobson et al., 2006). CRMP-2 links tubulin heterodimers or Sra-1 to Kinesin-1 through interaction with Kinesin light chain, and the CRMP-2/kinesin-1 complex regulates the transport of these proteins to the distal part of the growing axon (Kawano et al., 2005; Kimura et al., 2005). Thus CRMP-2 seems to promote neurite elongation and axon specification by regulating microtubule assembly, endocytosis of adhesion molecules, reorganization of actin filaments, and axonal protein trafficking (> Figure 3‐3). GSK-3b can phosphorylate CRMP-2 at Thr-514 and Ser-518 through the priming phosphorylation at Ser-522 by Cdk5 (Yoshimura et al., 2005). Phosphorylation of CRMP-2 by GSK-3b lowers CRMP-2’s activity for the interaction with tubulin and Numb (Arimura et al., 2005; Yoshimura et al., 2005). In cultured hippocampal neurons, about 30% of CRMP-2 is constitutively phosphorylated at Thr-514, and this phosphorylation is decreased by GSK-3 inhibitors. GSK-3b is activated only in the tips of axons in stage 3 neurons (Jiang et al., 2005). CRMP-2 phosphorylated at Thr-514 is enriched in the distal part of the growing axons but clearly not at the axonal growth cones, suggesting that there is a nonphosphorylated CRMP-2 pool at Thr-514 in the growing axonal growth cone (Yoshimura et al., 2005). Expression

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of constitutively active GSK-3b impaired neuronal polarization, whereas the nonphosphorylated form of CRMP-2 counteracted the inhibitory effects of GSK-3b, which indicates that GSK-3b regulates neuronal polarity through phosphorylation of CRMP-2 (Yoshimura et al., 2005). Zhou et al. (2004) reported that nerve growth factor-induced axon elongation is mediated by the PI3kinase/ILK/GSK-3b/adenomatous polyposis coli (APC) pathway in dorsal root ganglion neurons. GSK-3b phosphorylates MAP1B and APC (Goold et al., 1999; Zumbrunn et al., 2001; Trivedi et al., 2005). Phosphorylation of MAP1B by GSK-3b suppresses detyrosination of microtubules and decreases the number of stable microtubules. Binding of APC to microtubules increases microtubule stability, whereas phosphorylation of APC decreases interaction with microtubules. Hippocampal neurons derived from double-knockout mice with disrupted tau and MAP1B genes, which have redundant functions, show a defect in axon formation at stage 3 (Takei et al., 2000). These results suggest that GSK-3b regulates microtubule dynamics through microtubule-associating molecules and thereby governs neuronal polarity (> Figure 3‐3).

4.3 Positive Feedback Loop The prototypic Par genes were identified in C. elegans for their roles in directing asymmetric cell division during early development (Cowan and Hyman, 2004). The Par complex (including Par3, Par6, and aPKC) functions in various cell-polarization events, including axon specification (Ohno, 2001; Shi et al., 2003; Macara, 2004; Nishimura et al., 2004). Rho family small GTPases are major regulators of actin filaments and microtubules (Govek et al., 2005). Rho family small GTPases cycle between a GTP-bound active state and a GDP-bound inactive state, acting as molecular switches. Guanine nucleotide exchange factors (GEFs) activate GTPases by enhancing the exchange of bound GDP for GTP (Schmidt and Hall, 2002). Of the Rho family small GTPases, Cdc42, Rac1, and RhoA have been characterized most extensively. PI3-kinase activity is also required for proper localization of the Par complex and Cdc42 at the tips of the growing axons, both of which are necessary for neuronal polarization (Shi et al., 2003; Schwamborn and Puschel, 2004). Nishimura et al. (2004) reported that Par3 is transported to the distal tip of the growing axon by Kineisn-2 through direct interaction with KIF3A and that proper localization of Par3 is required to establish neuronal polarity. T-lymphoma invasion and metastasis 1 (Tiam1, a GEF for Rac1) are involved in axon formation (Kunda et al., 2001). Cdc42-GTP binds to Par6 and determines the localization of the Par complex. Par3 directly interacts with Sif- and Tiam1-like exchange factor (STEF)/Tiam1 (GEFs for Rac1), and the Par3/Par6 complex mediates the signal from Cdc42 to Rac1 for axon specification (Nishimura et al., 2005). Given that Rac1 activates PI3-kinase, the signal initially evoked by PI3-kinase appears to terminate at PI3-kinase itself (Govek et al., 2005). This positive feedback loop may be a driving force for axon specification and maturation (> Figure 3‐3). Recently, it was reported that the Rac activator DOCK7 is important for axon formation (WatabeUchida et al., 2006). DOCK7 is asymmetrically distributed in stage 2 hippocampal neurons and selectively expressed in the axon. Knockdown of DOCK7 prevents axon formation, whereas overexpression induces the formation of multiple axons. DOCK7 regulates Rac activity to inactivate the microtubule destabilizing protein stathmin/Op18 and promote axon formation (Watabe-Uchida et al., 2006). Localized activation of the p21-activated kinase is pivotal for neuronal polarity by affecting the function of proteins that regulate the actin filaments and microtubules through the activation of Rac (Jacobs et al., 2007). Thus DOCK7 and Tiam1 or STEF trigger the activation of Rac to regulate microtubule and actin remodeling, respectively.

4.4 Rap1B, H-Ras, and R-Ras It has been reported that Rap1B, H-Ras, and R-Ras, members of the Ras family small GTPases, are involved in axon specification (Schwamborn and Puschel, 2004; Yoshimura et al., 2006; Oinuma et al., 2007). Rap1B acts upstream of Cdc42 and the Par complex in neuronal polarity (Schwamborn and Puschel, 2004). Overexpression of Rap1B induces multiple axons and accumulation of the Par complex in each neurite.

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Knockdown of Rap1B causes the complete loss of axons. Analysis using inhibitors revealed that PI3-kinase functions upstream of Rap1B. What is the upstream signaling molecule of PI3-kinase on neuronal polarity? H-Ras and R-Ras play critical roles in establishing neuronal polarity upstream of the PI3-kinase/Akt/ GSK-3b pathway (Yoshimura et al., 2006; Oinuma et al., 2007). Ras may stimulate and activate PI3-kinase at the tip of one of the immature neurites downstream of the extracellular cues (> Figure 3‐3).

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Conclusion and Perspectives

Significant progress has been made toward understanding the intracellular signaling cascades during neuronal polarization. > Figure 3‐3 is a model schema of the signaling cascades in axon formation. Two major signaling cascades downstream of PI3-kinase play a central role in neuronal polarization. The PI3-kinase/Akt/GSK-3b/ CRMP-2 pathway promotes neurite outgrowth to determine axon or dendrite fate. The positive feedback loop composed of Cdc42, the Par complex, and Rac1 cycles locally in the tips of future axons. However, more questions must be answered before the molecular mechanisms will be entirely understood. It is conceivable that in vivo, extracellular cues play a pivotal role in the specification of axonal fate. What are the extracellular signals involved in the establishment of neuronal polarity? Axons and dendrites face the preferred direction in vivo, where extrinsic signals from the surrounding cellular environment likely play a major role in neuronal polarization. Little is known about extracellular cues that govern neuronal polarity in vivo. Additional studies are needed to elucidate fully the mechanisms behind establishment of neuronal polarity.

References Andersen SS, Bi GQ. 2000. Axon formation: A molecular model for the generation of neuronal polarity. Bioessays 22: 172-179. Arimura N, Menager C, Fukata Y, Kaibuchi K. 2004. Role of CRMP-2 in neuronal polarity. J Neurobiol 58: 34-47. Arimura N, Menager C, Kawano Y, Yoshimura T, Kawabata S, et al. 2005. Phosphorylation by Rho kinase regulates CRMP2 activity in growth cones. Mol Cell Biol 25: 9973-9984. Arimura N, Kaibuchi K. 2007. Neuronal polarity: From extracellular signals to intracellular mechanisms. Nat Rev Neurosci 8: 194-205. Baas PW. 1997. Microtubules and axonal growth. Curr Opin Cell Biol 9: 29-36. Baas PW, Buster DW. 2004. Slow axonal transport and the genesis of neuronal morphology. J Neurobiol 58: 3-17. Bradke F, Dotti CG. 1999. The role of local actin instability in axon formation. Science 283: 1931-1934. Bradke F, Dotti CG. 2000a. Establishment of neuronal polarity: Lessons from cultured hippocampal neurons. Curr Opin Neurobiol 10: 574-581. Bradke F, Dotti CG. 2000b. Differentiated neurons retain the capacity to generate axons from dendrites. Curr Biol 10: 1467-1470. Brown A, Slaughter T, Black MM. 1992. Newly assembled microtubules are concentrated in the proximal and distal regions of growing axons. J Cell Biol 119: 867-882. Brown A, Li Y, Slaughter T, Black MM. 1993. Composite microtubules of the axon: Quantitative analysis of

tyrosinated and acetylated tubulin along individual axonal microtubules. J Cell Sci 104: 339-352. Cowan CR, Hyman AA. 2004. Asymmetric cell division in C. elegans: Cortical polarity and spindle positioning. Annu Rev Cell Dev Biol 20: 427-453. Craig AM, Banker G. 1994. Neuronal polarity. Annu Rev Neurosci 17: 267-310. Dotti CG, Banker GA. 1987. Experimentally induced alteration in the polarity of developing neurons. Nature 330: 254-256. Dotti CG, Sullivan CA, Banker GA. 1988. The establishment of polarity by hippocampal neurons in culture. J Neurosci 8: 1454-1468. Esch T, Lemmon V, Banker G. 1999. Local presentation of substrate molecules directs axon specification by cultured hippocampal neurons. J Neurosci 19: 6417-6426. Forscher P, Smith SJ. 1988. Actions of cytochalasins on the organization of actin filaments and microtubules in a neuronal growth cone. J Cell Biol 107: 1505-1516. Fukata Y, Itoh TJ, Kimura T, Menager C, Nishimura T, et al. 2002. CRMP-2 binds to tubulin heterodimers to promote microtubule assembly. Nat Cell Biol 4: 583-591. Goold RG, Owen R, Gordon-Weeks PR. 1999. Glycogen synthase kinase 3b phosphorylation of microtubule-associated protein 1B regulates the stability of microtubules in growth cones. J Cell Sci 112: 3373-3384. Goshima Y, Nakamura F, Strittmatter P, Strittmatter SM. 1995. Collapsin-induced growth cone collapse mediated

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phosphatidylinositol 3,4,5-trisphosphate. J Biol Chem 273: 13375-13378. Menager C, Arimura N, Fukata Y, Kaibuchi K. 2004. PIP3 is involved in neuronal polarization and axon formation. J Neurochem 89: 109-118. Nakata T, Hirokawa N. 2003. Microtubules provide directional cues for polarized axonal transport through interaction with kinesin motor head. J Cell Biol 162: 1045-1055. Nishimura T, Fukata Y, Kato K, Yamaguchi T, Matsuura Y, et al. 2003. CRMP-2 regulates polarized Numb-mediated endocytosis for axon growth. Nat Cell Biol 5: 819-826. Nishimura T, Kato K, Yamaguchi T, Fukata Y, Ohno S, et al. 2004. Role of the PAR-3-KIF3 complex in the establishment of neuronal polarity. Nat Cell Biol 6: 328-334. Nishimura T, Yamaguchi T, Kato K, Yoshizawa M, Nabeshima Y, et al. 2005. PAR-6-PAR-3 mediates Cdc42induced Rac activation through the Rac GEFs STEF/Tiam1. Nat Cell Biol 7: 270-277. Ohno S. 2001. Intercellular junctions and cellular polarity: The PAR–aPKC complex, a conserved core cassette playing fundamental roles in cell polarity. Curr Opin Cell Biol 13: 641-648. Oinuma I, Katoh H, Negishi M. 2007. R-Ras controls axon specification upstream of glycogen synthase kinase-3beta through integrin-linked kinase. J Biol Chem 282: 303-318. Scheid MP, Woodgett JR. 2001. PKB/AKT: Functional insights from genetic models. Nat Rev Mol Cell Biol 2: 760-768. Schmidt A, Hall A. 2002. Guanine nucleotide exchange factors for Rho GTPases: Turning on the switch. Genes Dev 16: 1587-1609. Schwamborn JC, Puschel AW. 2004. The sequential activity of the GTPases Rap1B and Cdc42 determines neuronal polarity. Nat Neurosci 7: 923-929. Shi SH, Jan LY, Jan YN. 2003. Hippocampal neuronal polarity specified by spatially localized mPar3/mPar6 and PI 3-kinase activity. Cell 112: 63-75. Takei Y, Teng J, Harada A, Hirokawa N. 2000. Defects in axonal elongation and neuronal migration in mice with disrupted tau and map1b genes. J Cell Biol 150: 989-1000. Toriyama M, Shimada T, Kim KB, Mitsuba M, Nomura E, et al. 2006. Shootin1: A protein involved in the organization of an asymmetric signal for neuronal polarization. J Cell Biol 175: 147-157. Trivedi N, Marsh P, Goold RG, Wood-Kaczmar A, GordonWeeks PR. 2005. Glycogen synthase kinase-3b phosphorylation of MAP1B at Ser1260 and Thr1265 is spatially restricted to growing axons. J Cell Sci 118: 993-1005. Wang LH, Strittmatter SM. 1996. A family of rat CRMP genes is differentially expressed in the nervous system. J Neurosci 16: 6197-6207.

Regulation of axon formation Watabe-Uchida M, John KA, Janas JA, Newey SE, Van Aelst L. 2006. The Rac activator DOCK7 regulates neuronal polarity through local phosphorylation of stathmin/Op18. Neuron 51: 727-739. Yan D, Guo L, Wang Y. 2006. Requirement of dendritic Akt degradation by the ubiquitin-proteasome system for neuronal polarity. J Cell Biol 174: 415-424. Yoshimura T, Kawano Y, Arimura N, Kawabata S, Kikuchi A, et al. 2005. GSK-3b regulates phosphorylation of CRMP-2 and neuronal polarity. Cell 120: 137-149. Yoshimura T, Arimura N, Kawano Y, Kawabata S, Wang S, et al. 2006. Ras regulates neuronal polarity via the PI3-kinase/Akt/GSK-3b/CRMP-2 pathway. Biochem Biophys Res Commun 340: 62-68.

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Neuronal Process Outgrowth

T. Mori . N. Inagaki . H. Kamiguchi

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40 2 Clutch Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40 3 Neurite Initiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41 4 Neurite Elongation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42 5 Neurite Differentiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42 6 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43

#

2009 Springer Science+Business Media, LLC.

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Abstract: Cell motility depends on the coordinated functions of the cytoskeleton and cell adhesion molecules. Their dynamic interactions are particularly important for directing polarized behavior of a cell. A neuron forms and elongates neurites that eventually become an axon or dendrites. These developmental processes also require coordinated activities of the cytoskeleton and cell adhesion molecules in a spatiotemporally regulated manner. In this chapter, we will summarize the molecular mechanisms of neurite growth with particular emphasis on a molecular clutch that mechanically links the cytoskeleton to the extracellular environment through cell adhesion molecules. List of Abbreviations: CAM, cell adhesion molecule; ECM, extracellular matrix; IgCAM, immunoglobulin superfamily CAM

1 Introduction During development, neurons extend morphologically and functionally differentiated processes, axons, and dendrites. Over a decade ago, Banker and colleagues developed a culture system of rodent embryonic hippocampal neurons and precisely observed the morphological changes of neurons during maturation (Dotti et al., 1988; Goslin and Banker, 1989). They divided the process of neurite formation and differentiation into five stages: first, a neuron generates lamellipodia surrounding the cell body (Stage1). Several hours later, the neuron extends multiple immature neurites called minor processes (Stage2), which are morphologically indistinguishable from each other. Within 24 h in culture, one of the minor processes elongates faster than the other processes and becomes an axon (Stage3). As the axon elongates, the other neurites gradually grow and become the dendrites (Stage4). More than 10–14 days later, the axon and the dendrites form the specific structure, called the synapse, and construct neural circuits (Stage5). In this chapter, we will summarize the recent progress of our understanding of neurite initiation, elongation, and polarization, with particular emphasis on cell adhesion molecules (CAMs) and the cytoskeleton.

2 Clutch Mechanism Spatially localized actin polymerization/depolymerization and actin–myosin interactions generate retrograde flow of filamentous actin (F-actin) in motile cells (Pollard and Borisy, 2003). In neuronal growth cones, myosin 1c and myosin II have been implicated in moving F-actin in the retrograde direction (Diefenbach et al., 2002; Medeiros et al., 2006). It is widely accepted that retrograde F-actin flow generates traction force that drives cell migration and process outgrowth. According to the clutch hypothesis (Mitchison and Kirschner, 1988), CAMs transmit this force by linking the F-actin flow with immobile ligands present on neighboring cells or in the extracellular matrix (ECM). A molecular clutch, which is composed of multiple molecules, mediates the engagement between the F-actin flow and CAMs in a spatiotemporally regulated manner. So far, three major classes of CAMs have been identified in the nervous system: integrins, cadherins, and the immunoglobulin superfamily CAMs (IgCAMs). Cadherins and the majority of IgCAMs mediate cell– cell adhesion via a homophilic binding mechanism, whereas integrins interact with ECM molecules. It has been shown that several IgCAMs, such as apCAM, NrCAM, and L1, couple with retrograde F-actin flow in neurons, thereby promoting neurite initiation or elongation. Two molecules, ankyrinB (Nishimura et al., 2003) and shootin1 (Shimada et al., 2006), have been identified as a component of the clutch module that connects L1 with F-actin flow. It has also been suggested that catenins mediate the linkage between N-cadherin and F-actin flow in neuronal growth cones (Thoumine et al., 2006). More recently, two important reports have been published that address the clutch mechanism in nonneuronal cells that migrate on ECM substrates using integrins as adhesive receptors (Brown et al., 2006; Hu et al., 2007). Both studies employed total internal reflection microscopy to analyze the dynamics of focal adhesion proteins with regard to F-actin and integrins. F-actin flowed retrogradely at the site of focal adhesions, whereas

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integrins in the same region remained largely stationary. Different classes of focal adhesion proteins exhibited intermediate and varying degrees of retrograde movement, indicating hierarchical transmission of F-actin flow through focal adhesions that constitute a slippage interface of the clutch module.

3 Neurite Initiation Halpain and colleagues investigated the process of neurite initiation and found that segmented lamellipodia are precursors of neurites in hippocampal neurons in culture (Dehmelt et al., 2003; Dehmelt and Halpain, 2004): Perisomatic lamellipodia, which uniformly surround the cell body, become segmented at one or more sites. These segmented areas accumulate microtubules and gradually migrate away from the cell body, transforming the lamellipodia into growth cones. Then, the newly formed protrusions elongate and the microtubules become tightly packed inside the neurite shafts. How does a neuron determine the active regions of lamellipodia that eventually become neurites? It has been suggested that the linkage of F-actin with extracellular substrates via CAMs is involved in the specification of neurite-forming areas within the perisomatic lamellipodia (Nishimura et al., 2003). While it was hypothesized that the clutch mechanism is involved in the initial formation of neurites (Smith, 1994), the molecular identity of the clutch module was poorly understood. Using dorsal root ganglion neurons in culture, we showed that ankyrinB, a protein associating with the spectrin–actin network, constitutes the clutch module that regulates neurite initiation stimulated by L1 (Nishimura et al., 2003). In the perisomatic protrusions before neuritogenesis, ankyrinB exhibits retrograde (centripetal) movement that is dependent on F-actin flow toward the cell body. A mutant form of ankyrinB that has a single amino acid substitution in its spectrin-binding domain does not move retrogradely, suggesting that ankyrinB binds to F-actin flow via spectrin. By monitoring the behavior of a bead bound to L1 on the perisomatic lamellipodium, we showed that L1 couples with retrograde F-actin flow and this coupling is mediated, in part, by ankyrinB. Ligation of the L1 ectodomain by an immobile substrate induces L1-ankyrinB binding and the formation of stationary clusters of ankyrinB in the perisomatic protrusions. Neurite initiation preferentially occurs at the site of these clusters. Furthermore, neurite initiation is impaired when the L1 linkage with F-actin flow is inhibited by ankyrinB knockout. These results support the idea that ankyrinB promotes neurite initiation by acting as a component of the clutch module that transmits traction force generated by F-actin flow to the extracellular substrate via L1. In contrast, ankyrinB is involved neither in L1 coupling with F-actin flow in growth cones nor in L1-mediated neurite elongation, indicating that neurons change components of the clutch module as they mature and that ankyrinB functions as a clutch only before neurite formation. Some form of ankyrin acts even as a negative regulator of L1 coupling with F-actin flow in a growth cone-like structure of ND-7 neuroblastoma hybrid cells (Gil et al., 2003). For the continued protrusion of the active region of perisomatic lamellipodia, membrane components together with functional molecules must be supplied to the tip of emerging neurites. Microtubules play a central role in the transport of organelles and molecules. However, microtubules do not reach the leading edge of lamellipodia before neuritogenesis, most likely because retrograde F-actin flow prevents microtubules from invading the cell periphery (Rodriguez et al., 2003). While F-actin functions as guides along which microtubules grow toward the leading edge, retrograde F-actin flow moves microtubules rearward, often resulting in breakage and subsequent turnover of the microtubules (Gupton et al., 2002; Salmon et al., 2002; Schaefer et al., 2002a). When the flow rate of F-actin in the perisomatic lamellipodia attenuates as a result of the linkage with extracellular immobile substrates, microtubules grow into the area of clutch engagement leading to the formation of neurites. In turn, the microtubule growth may promote lamellipodial protrusion by activating Rac, a Rho-family GTPase that promotes the formation of actin network in the lamellipodia (Waterman-Storer et al., 1999). It is also likely that active Rac stabilizes microtubules by inhibiting the microtubule-destabilizing protein OP180/Stathmin (Daub et al., 2001), forming a positivefeedback loop that dramatically reorganizes both microtubules and F-actin during neuritogenesis. In this way, local extracellular environment may determine the site of neuritogenesis by binding CAMs and engaging a molecular clutch, which eventually influences the dynamics and organization of the cytoskeleton in a manner important for extending neurites.

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4 Neurite Elongation The clutch mechanism also plays a critical role in transmitting traction force for neurite elongation, and shootin1 has been implicated as a component of the clutch module that links L1 with retrograde F-actin flow in the growth cone (Shimada et al., 2006). As the growth cone migrates forward, L1 translocates rearward into the central domain of the growth cone and is internalized via clathrin-dependent pathways (Kamiguchi et al., 1998). The phosphorylation state of a tyrosine residue in the L1 cytosplasmic domain regulates L1 endocytosis by altering its binding affinity to the clathrin adaptor AP2 (Schaefer et al., 2002b). Subsequently, internalized L1 is transported toward the growth cone periphery along microtubules and recycled to the plasma membrane of the leading front (Kamiguchi and Lemmon, 2000). The intracellular transport of L1-containing vesicles may be driven by KIF4, a plus-end-directed motor along a microtubule (Peretti et al., 2000). L1 exocytosis, as a result of vesicle fusion with the plasma membrane, may be mediated by tetanus neurotoxin-insensitive vesicle-associated membrane protein (Alberts et al., 2003). In this way, forward migration of the growth cone is driven by spatially regulated endocytosis and exocytosis together with cell surface movement of L1 (Kamiguchi and Yoshihara, 2001). Recently, Thoumine and colleagues conducted a series of detailed experiments and have demonstrated the similar type of L1 trafficking in growth cones (Dequidt et al., 2007). Another example of CAM trafficking that drives neurite elongation came from studies on integrins. Integrin-dependent adhesion sites in growth cones, known as point contacts, are critically involved in neurite elongation (Robles et al., 2005; Woo and Gomez, 2006). It has been reported that b1 integrin exhibits bidirectional movement on the growth cone surface: substratebound integrin moves rearward, whereas unbound integrin is recycled to the leading front (Grabham and Goldberg, 1997; Grabham et al., 2000). Therefore, the bidirectional movement of CAMs may be a general mechanism for growth cone migration although distinct recycling pathways and diverse clutch molecules may exist for trafficking of different CAMs.

5 Neurite Differentiation The initial event in neuronal polarization is the specification of an axon. It has been reported that the neurite which is over 10 mm longer than the other minor processes becomes an axon, implying that the length of neurites is an important factor for axon specification (Goslin and Banker, 1989). When the growth cone of one neurite contacts growth-promoting substrate, that neurite becomes an axon (Esch et al., 1999). Therefore, extracellular environments seem to provide a signal that is sufficient for axon specification. Bradke and Dotti (1999) reported that the tip of the future axon contains less dense network of F-actin than the tip of the other minor processes and that a treatment of stage-2 neurons with cytochalasin D, an actin-depolymerizing drug, results in the formation of multiple axons. The F-actin network can prevent the microtubule from invading the growth cone periphery and inhibit neurite elongation. Therefore, local instability of the F-actin network restricted to a single growth cone allows the neurite to elongate faster and may act as a physiological signal for axon specification. Now it is widely accepted that establishment of axon–dendrite polarity depends on various molecules involved in signal transduction, cytoskeletal organization, vesicle transport, lipid metabolism, and protein degradation (Arimura and Kaibuchi, 2005; Da Silva et al., 2005; Yan et al., 2006). However, many molecules carrying out this task remain to be identified. To search for such molecules, Inagaki and colleagues performed proteomic analyses and compared the expression profiles of proteins between stage 2 and stage 3 hippocampal neurons in culture (Toriyama et al., 2006). It was found that, using a large gel (93  103 cm) for highresolution two-dimensional electrophoresis, 277 out of 6,000 protein spots showed consistent upregulation during the transition from stage 2 to stage 3. So far, two novel proteins, shootin1 and singar1, have been identified and reported (Toriyama et al., 2006; Mori et al., 2007). Shootin1 is expressed predominantly in axonal growth cones of embryonic and early postnatal neurons. Overexpression of shootin1 induces the formation of multiple axons, and suppression of shootin1 causes a delay in axon formation, suggesting that shootin1 is a positive regulator for axon formation perhaps by acting as a molecular clutch. Singar1 and its

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splicing variant, singar2, are expressed in both an axon and somatodendrites. Although overexpression of singar1 and singar2 does not influence the number of axons per neuron, suppression of both singar1 and singar2 induces the formation of multiple axons. Interestingly, overexpression of singar1, but not singar2, reduces the number of neurons bearing surplus axons induced by excessive amounts of shootin1. Although the functional difference between singar1 and singar2 is unclear, the data indicate that singar1 suppresses the formation of surplus axons without affecting normal polarization processes thereby contributing to the robustness of axon–dendrite polarity.

6 Conclusions Components of the clutch module that regulate neurite initiation and elongation have just begun to be identified. Furthermore, mechanisms of slipping engagement of the clutch module have been another subject of recent studies. Because neurons employ distinct sets of clutch components depending not only on CAMs used to interact with extracellular environments but also on developmental stages of neurites, a number of clutch molecules and mechanisms controlling their engagement remain to be investigated. Future studies of these problems will provide a clear picture of how the dynamic and regulated interactions between CAMs and the cytoskeleton promote neurite growth and control axon specification.

References Alberts P, Rudge R, Hinners I, Muzerelle A, Martinez-Arca S, et al. 2003. Cross talk between tetanus neurotoxin-insensitive vesicle-associated membrane protein-mediated transport and L1-mediated adhesion. Mol Biol Cell 14: 4207-4220. Arimura N, Kaibuchi K. 2005. Key regulators in neuronal polarity. Neuron 48: 881-884. Bradke F, Dotti CG. 1999. The role of local actin instability in axon formation. Science 283: 1931-1934. Brown CM, Hebert B, Kolin DL, Zareno J, Whitmore L, et al. 2006. Probing the integrin-actin linkage using highresolution protein velocity mapping. J Cell Sci 119: 5204-5214. Da Silva JS, Hasegawa T, Miyagi T, Dotti CG, Abad-Rodriguez J. 2005. Asymmetric membrane ganglioside sialidase activity specifies axonal fate. Nat Neurosci 8: 606-615. Daub H, Gevaert K, Vandekerckhove J, Sobel A, Hall A. 2001. Rac/Cdc42 and p65PAK regulate the microtubuledestabilizing protein stathmin through phosphorylation at serine 16. J Biol Chem 276: 1677-1680. Dehmelt L, Halpain S. 2004. Actin and microtubules in neurite initiation: Are MAPs the missing link? J Neurobiol 58: 18-33. Dehmelt L, Smart FM, Ozer RS, Halpain S. 2003. The role of microtubule-associated protein 2c in the reorganization of microtubules and lamellipodia during neurite initiation. J Neurosci 23: 9479-9490. Dequidt C, Danglot L, Alberts P, Galli T, Choquet D, et al. 2007. Fast turnover of L1 adhesions in neuronal growth cones involving both surface diffusion and exo/endocytosis of L1 molecules. Mol Biol Cell 18: 3131-3143.

Diefenbach TJ, Latham VM, Yimlamai D, Liu CA, Herman IM, et al. 2002. Myosin 1c and myosin IIB serve opposing roles in lamellipodial dynamics of the neuronal growth cone. J Cell Biol 158: 1207-1217. Dotti CG, Sullivan CA, Banker GA. 1988. The establishment of polarity by hippocampal neurons in culture. J Neurosci 8: 1454-1468. Esch T, Lemmon V, Banker G. 1999. Local presentation of substrate molecules directs axon specification by cultured hippocampal neurons. J Neurosci 19: 6417-6426. Gil OD, Sakurai T, Bradley AE, Fink MY, Cassella MR, et al. 2003. Ankyrin binding mediates L1CAM interactions with static components of the cytoskeleton and inhibits retrograde movement of L1CAM on the cell surface. J Cell Biol 162: 719-730. Goslin K, Banker G. 1989. Experimental observations on the development of polarity by hippocampal neurons in culture. J Cell Biol 108: 1507-1516. Grabham PW, Foley M, Umeojiako A, Goldberg DJ. 2000. Nerve growth factor stimulates coupling of b1 integrin to distinct transport mechanisms in the filopodia of growth cones. J Cell Sci 113: 3003-3012. Grabham PW, Goldberg DJ. 1997. Nerve growth factor stimulates the accumulation of b1 integrin at the tips of filopodia in the growth cones of sympathetic neurons. J Neurosci 17: 5455-5465. Gupton SL, Salmon WC, Waterman-Storer CM. 2002. Converging populations of f-actin promote breakage of associated microtubules to spatially regulate microtubule turnover in migrating cells. Curr Biol 12: 1891-1899.

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Hu K, Ji L, Applegate KT, Danuser G, Waterman-Storer CM. 2007. Differential transmission of actin motion within focal adhesions. Science 315: 111-115. Kamiguchi H, Lemmon V. 2000. Recycling of the cell adhesion molecule L1 in axonal growth cones. J Neurosci 20: 3676-3686. Kamiguchi H, Long KE, Pendergast M, Schaefer AW, Rapoport I, et al. 1998. The neural cell adhesion molecule L1 interacts with the AP-2 adaptor and is endocytosed via the clathrin-mediated pathway. J Neurosci 18: 5311-5321. Kamiguchi H, Yoshihara F. 2001. The role of endocytic L1 trafficking in polarized adhesion and migration of nerve growth cones. J Neurosci 21: 9194-9203. Medeiros NA, Burnette DT, Forscher P. 2006. Myosin II functions in actin-bundle turnover in neuronal growth cones. Nat Cell Biol 8: 215-226. Mitchison T, Kirschner M. 1988. Cytoskeletal dynamics and nerve growth. Neuron 1: 761-772. Mori T, Wada T, Suzuki T, Kubota Y, Inagaki N. 2007. Singar1, a novel RUN domain-containing protein, suppresses formation of surplus axons for neuronal polarity. J Biol Chem 282: 19884-19893. Nishimura K, Yoshihara F, Tojima T, Ooashi N, Yoon W, et al. 2003. L1-dependent neuritogenesis involves ankyrinB that mediates L1-CAM coupling with retrograde actin flow. J Cell Biol 163: 1077-1088. Peretti D, Peris L, Rosso S, Quiroga S, Caceres A. 2000. Evidence for the involvement of KIF4 in the anterograde transport of L1-containing vesicles. J Cell Biol 149: 141-152. Pollard TD, Borisy GG. 2003. Cellular motility driven by assembly and disassembly of actin filaments. Cell 112: 453-465. Robles E, Woo S, Gomez TM. 2005. Src-dependent tyrosine phosphorylation at the tips of growth cone filopodia promotes extension. J Neurosci 25: 7669-7681. Rodriguez OC, Schaefer AW, Mandato CA, Forscher P, Bement WM, et al. 2003. Conserved microtubule-actin interactions in cell movement and morphogenesis. Nat Cell Biol 5: 599-609.

Salmon WC, Adams MC, Waterman-Storer CM. 2002. Dualwavelength fluorescent speckle microscopy reveals coupling of microtubule and actin movements in migrating cells. J Cell Biol 158: 31-37. Schaefer AW, Kabir N, Forscher P. 2002a. Filopodia and actin arcs guide the assembly and transport of two populations of microtubules with unique dynamic parameters in neuronal growth cones. J Cell Biol 158: 139-152. Schaefer AW, Kamei Y, Kamiguchi H, Wong EV, Rapoport I, et al. 2002b. L1 endocytosis is controlled by a phosphorylation-dephosphorylation cycle stimulated by outside-in signaling by L1. J Cell Biol 157: 1223-1232. Shimada T, Toriyama M, Uemura K, Sugiura T, Watanabe N, et al. 2006. Shootin1 transmits driving force of F-actin flow to L1-CAM for neurite elongation. 2006 Cold Spring Harbor Meeting Abstract: 177. Smith CL. 1994. Cytoskeletal movements and substrate interactions during initiation of neurite outgrowth by sympathetic neurons New York: in vitro. J Neurosci 14: 384-398. Thoumine O, Lambert M, Mege RM, Choquet D. 2006. Regulation of N-cadherin dynamics at neuronal contacts by ligand binding and cytoskeletal coupling. Mol Biol Cell 17: 862-875. Toriyama M, Shimada T, Kim KB, Mitsuba M, Nomura E, et al. 2006. Shootin1: A protein involved in the organization of an asymmetric signal for neuronal polarization. J Cell Biol 175: 147-157. Waterman-Storer CM, Worthylake RA, Liu BP, Burridge K, Salmon ED. 1999. Microtubule growth activates Rac1 to promote lamellipodial protrusion in fibroblasts. Nat Cell Biol 1: 45-50. Woo S, Gomez TM. 2006. Rac1 and RhoA promote neurite outgrowth through formation and stabilization of growth cone point contacts. J Neurosci 26: 1418-1428. Yan D, Guo L, Wang Y. 2006. Requirement of dendritic Akt degradation by the ubiquitin-proteasome system for neuronal polarity. J Cell Biol 174: 415-424.

Signaling in Synaptic Transmission

5

Proteins Involved in the Presynaptic Functions

M. Igarashi . K. Ohko

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48

2 2.1 2.2 2.2.1 2.2.2 2.2.3 2.3 2.3.1 2.3.2 2.3.3 2.3.4

The SNARE Complex: The Main Molecular Machinery for Ca2+-Regulated Exocytosis . . . . . . The Concept of the SNARE Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . SNAREs and Related Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Syntaxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . SNAP-25 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VAMP-2 (synaptobrevin) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulators of the SNARE Proteins or the SNARE Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . NSF (N-ethylmaleimide-sensitive factor) and SNAPs (soluble NSF-associated proteins) . . . . . . . . . . . . Munc-18 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Complexin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tomosyn . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

48 48 48 48 49 50 50 50 50 50 51

3 3.1 3.1.1 3.1.2 3.1.3 3.1.4 3.1.5 3.1.6 3.2 3.2.1 3.2.2 3.2.3 3.2.4 3.3 3.3.1 3.3.2

Catalog of the Presynaptic Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Other SV Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Synaptophysin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Synaptic Glycoprotein 2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Synapsins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rab3A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vacuolar-ATPase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cysteine String Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Active Zone Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Munc13 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Bassoon and Piccolo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . CAST/ELKs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . RIM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Presynaptic Membrane Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Neurexins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ca2+ Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

51 51 51 52 52 52 52 52 53 53 54 54 54 54 54 54

The Proteins Involved in Ca2+-Dependent Vesicular Recycling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Synaptotagmins and Low-Affinity Ca2+ Sensors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ca2+-Dependent Vesicular Recycling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Submicromolar Ca2+ Concentrations Regulate Vesicular Recycling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . CaM-Dependent Regulation of Exocytosis Through Modulation of Membrane Trafficking Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.1 CaMKII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.2 Myosin-V . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4.3 VAMP/Synaptobrevin and CaM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

55 55 55 55

4 4.1 4.2 4.3 4.4

5 #

56 56 57 59

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 2009 Springer Science+Business Media, LLC.

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Abstract: The proteins specific to the presynaptic terminal have been well-characterized, and a large number of the interactions among these proteins have been elucidated. The roles of some of these proteins in neurotransmitter release and synaptic vesicle recycling can be determined from their interactions, but electrophysiological data suggest that there remain additional steps whose molecular mechanisms are not yet known. In this review, we summarize the characteristics of each presynaptic protein, and we discuss some of the important interactions, including their regulation by Ca2+ and how they participate in the Ca2+-dependent steps of vesicular recycling. List of Abbreviations: CaMKII, calmodulin kinase II; CSP, cysteine string protein; NSF, N-ethylmaleimide-sensitive factor; RRP, rapidly releasable pool; SNAPs, soluble NSF-associated proteins; t-SNARE, target SNARE; V-ATPase, vacuolar-ATPase; v-SNARE, vesicular SNARE; VAMP-2, vesicle-associated membrane protein-2

1

Introduction

Presynaptic proteins are involved in each step of neurotransmitter release. In the last two decades, more than 20 species of presynaptic proteins have been biochemically and functionally characterized. This information has helped clarify the fundamental molecular machinery of exocytosis and vesicular recycling. For some of these proteins, their presynaptic functions are not completely known, but there are many interactions between them, suggesting that they participate in vesicular recycling. In this review, we describe the characteristics of the presynaptic proteins and their roles in exocytosis and vesicular recycling. In particular, we discuss several protein–protein interactions that are dependent on Ca2+ and are involved in the Ca2+-dependent steps of exocytosis and vesicular recycling.

2

The SNARE Complex: The Main Molecular Machinery for Ca2+-Regulated Exocytosis

2.1 The Concept of the SNARE Mechanism The SNARE mechanism is considered the main molecular mediator of the interaction between synaptic vesicles (SVs) and the plasma membrane (> Figure 5‐1a; Duman and Forte, 2003). Its key concept is that a vesicular SNARE (v-SNARE) protein on the vesicle and a target SNARE (t-SNARE) protein on the target membrane form a complex (i.e., the SNARE complex) that mediates the interaction between the vesicle and the target (Li and Chin, 2003). VAMP-2 (vesicle-associated membrane protein-2)/synaptobrevin is a v-SNARE on SVs, and SNAP-25 (synaptosomal-associated protein 25 kDa) and syntaxin-1A act as t-SNAREs in the presynaptic membrane. This process probably corresponds to the docking of the SVs (> Figure 5‐1a and > 1b). In addition, the SNARE complex itself is able to stimulate vesicular fusion.

2.2 SNAREs and Related Proteins 2.2.1 Syntaxin The syntaxins comprise the largest protein family involved in vesicular trafficking. Five syntaxins (1A, 1B, 2, 3, and 4) are localized in the plasma membrane (Bennett et al., 1993), and syntaxin-1A and -1B are specific to neurons and secretory cells such as chromaffin cells. Syntaxin-1A is expressed at a higher level than syntaxin-1B, and the localization of these two species in brain regions are different (Gurkan et al., 2005). Nevertheless, their molecular interactions are similar. Syntaxin-1A interacts with SNARE proteins, regulators of the SNARE complex, channels, and transporters; thus, syntaxin-1A is the most important protein for regulation of exocytosis. Syntaxin-1A has three N-terminal helices (HA, HB, and HC domains), a C-terminal helix (H3 domain), a linker domain connecting the helices, and a C-terminal transmembrane

Proteins involved in the presynaptic functions

5

. Figure 5‐1 (a) The SNARE mechanism. The SNARE complex is composed of syntaxin, SNAP-25, and VAMP. This complex mediates vesicle docking and fusion. NSF and a-SNAP then dissociate the SNARE complex, allowing the reuse of the SNARE proteins. (b) Structure of rat syntaxin-1A. Syntaxin-1A has three N-terminal helices (HA, HB, and HC domains, collectively referred to as Habc), a C-terminal helix (H3 domain) containing the SNARE motif, a linker domain connecting the Habc and H3 domains, and a C-terminal transmembrane domain (TMR). (c) Conformational shift of syntaxin-1A (Dulubova et al., 1999). There are two forms of syntaxin-1A. In the closed form, the H3 domain, which contains the SNARE-binding site, is folded with the linker domain (residues 145–190) and the three N-terminal helices so that it cannot form a SNARE complex. In contrast, in the open form, the H3 domain is liberated from the linker domain and N-terminal helices, allowing formation of the SNARE complex. The conversion between these two forms is extremely rapid (0.8 ms; Margittai et al., 2003)

domain (> Figure 5‐1b; Dulubova et al., 1999). Syntaxin-1A exists in a closed and an open form (> Figure 5‐1c). Although the H3 domain serves as the binding site for other SNARE proteins, in the closed form, this site is buried in the interior of the protein by the N-terminal helices and cannot bind other SNARE proteins. In the open form, the H3 domain is exposed and can form the SNARE complex (Dulubova et al., 1999). Since the conversion between these two forms is very rapid (within 0.8 ms), it is likely that a stable SNARE complex formation is mediated by syntaxin-binding proteins fixing syntaxin in the open form (Margittai et al., 2003) and that the linker domain is the key site mediating this conformational change. A very recent report that employed far-field optical nanoscopy revealed that syntaxin molecules freely diffuse in the plasma membrane to form clusters of approximately 75 molecules, suggesting that the clustering of syntaxin is important in vesicle secretion (Sieber et al., 2007).

2.2.2 SNAP-25 The SNARE complex requires four a-helices, two of which are derived from syntaxin-1A and VAMP-2. The other two are derived from N- and C-terminus of SNAP-25, which is the other t-SNARE in the neuron

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besides syntaxin-1A. This protein lacks a transmembrane domain and associates with the plasma membrane via attached palmitoyl groups. There are two isoforms of SNAP-25 (SNAP-23 and -29), but the major component in the neuron is SNAP-25.

2.2.3 VAMP-2 (synaptobrevin) SVs have target-recognizing protein that recognizes its counterparts in the presynaptic membrane. These proteins are known as t-SNAREs. VAMP-2 is such a SV membrane protein. VAMP-2 has an a-helical cytoplasmic domain and a transmembrane domain.

2.3 Regulators of the SNARE Proteins or the SNARE Complex 2.3.1 NSF (N-ethylmaleimide-sensitive factor) and SNAPs (soluble NSF-associated proteins) NSF is an atypical ATPase protein, belongs to the AAA ATPase family, and is inhibited by N-ethylmaleimide. Although there are some different hypotheses regarding the role of NSF, currently, NSF is considered a molecular chaperone that disrupts the SNARE complex, enabling each SNARE protein to be reused. NSF interacts with SNAPs, such as a/b/g-SNAP, which mediate association with the SNAP receptors (also known as SNAREs) (Neuwald, 1999). NSF and SNAPs are not only localized in the presynaptic terminals but also in the Golgi apparatus and in other compartments. A variety of studies suggest that NSF acts as a chaperone for the SNARE proteins via the SNAPs. Specifically, a hexamer of NSF attaches to a trimer of SNAPs and then binds to the SNARE complex. In vitro studies have shown that this large complex dissociates the SNARE complex into free SNAREs in an ATP-dependent manner (Neuwald, 1999). This process is believed to allow the reuse of SNARE proteinsafter exocytosis (Rizo and Sudhof, 2002). NSF is also known to bind glutamate receptors in the postsynaptic area, acting as chaperone for their recycling (Osten et al., 1998).

2.3.2 Munc-18 Munc-18 is a mammalian homolog of Caenorhabditis elegans unc-18 and of Sec1p in Saccharomyces cerevisiae (budding yeast), which form the SM protein family. Although these family molecules do not have the transmembrane domain, these are mainly associated with the plasma membrane. Munc-18-1, a neuronal isoform of Munc-18, is tightly bound to syntaxin-1A. This interaction was originally interpreted as evidence that Munc-18-1 acts as inhibitor of exocytosis by sequestering syntaxin from the SNARE complex (Hata et al., 1993). However, this hypothesis was found to be incorrect because the exocytotic activity is not enhanced but rather inhibited in excitatory neurons from Munc-18-1 knockout mice (Verhage et al., 2000). This concept has been recently replaced by the idea that Munc-18 is not only bound to free syntaxin but also to syntaxin with SNARE complexes (Shen et al., 2007). In this way, Munc-18 can interact with the SNARE complex, thereby enhancing vesicular fusion, so that it is likely that this protein also participates in docking and priming by regulating syntaxin (Gulyas-Kovacs et al., 2007). Recent studies using chromaffin cells from the Munc-18-1 knockout mice revealed that Munc18-1 may participate in vesicular tethering via F-actin (Toonen et al., 2006). Thus, Munc-18 acts at several steps of vesicular recycling via a variety of interactions.

2.3.3 Complexin Complexin (also known as synaphin) is a small protein that binds to the SNARE complex at a 1:1 stoichiometry in a Ca2+-independent manner (Tokumaru et al., 2001). Mice lacking complexin I and II (the two are highly homologous) show a reduction in neurotransmitter release, which is thought to be due to a conformational change in the SNARE complex (Tang et al., 2006).

Proteins involved in the presynaptic functions

5

2.3.4 Tomosyn Tomosyn is a large, soluble, brain-specific syntaxin-binding protein. It competes with VAMP, preventing the formation of the SNARE complex. Thus, this protein is also a negative regulator of the SNARE complex (Fujita et al., 1998).

3

Catalog of the Presynaptic Proteins

3.1 Other SV Proteins Because SVs carry neurotransmitters, they contain the proteins for neurotransmitter condensation and for SV trafficking (> Figure 5‐2). The latter are independent of the specific neurotransmitters stored. A recently published study used a proteomic approach to determine the precise stoichiometry of the SV proteins (Takamori et al., 2006). Not all of the functions of the SV-specific proteins are defined, a number of their molecular interactions are known, which should help clarify their functions.

3.1.1 Synaptophysin Synaptophysin is a SV membrane protein with four transmembrane domains. This protein is widely accepted as the most reliable SV marker, but its function is uncertain. In vitro studies show that

. Figure 5‐2 Synaptic vesicle (SV) proteins. The major SV proteins are shown, although they are not shown in proportion to their known stoichiometries

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this protein interacts with VAMP-2 to prevent it from forming the SNARE complex (Edelmann et al., 1995). Knockout mice lacking synaptophysin show only a subtle electrophysiological change (McMahon et al., 1996).

3.1.2 Synaptic Glycoprotein 2 Synaptic glycoprotein 2 (SV2) is a SV membrane glycoprotein with 12 transmembrane domains, similar to membrane transporters; however, SV2 has not been shown to act as a transporter. SV2 has three homologs (SV2A, SV2B, and SV2C). SV2B interacts with synaptotagmin (Lazzell et al., 2004), but knockout mice lacking SV2A or SV2B show relatively a subtle electrophysiological change, and studies of exocytosis in cells from these mice has provided conflicting information on whether SV2 is involved in Ca2+ regulation (Janz et al., 1999; Custer et al., 2006).

3.1.3 Synapsins Synapsins were the first characterized presynaptic proteins. The synapsin family consists of synapsin Ia/b, synapsin IIa/b, and synapsin III. Synapsins undergo phosphorylation by calmodulin kinase II (CaMKII) and bind to SVs via phospholipids (Ferreira and Rapoport, 2002). Knockout mice lacking some of the synapsins, including double- or triple-knockout mice, do not show any phenotypes or any electrophysiological abnormalities consistent with the classical hypothesis for the role of synapsins. Thus, synapsin in the presynaptic terminal are not included in the minimal requirement of exocytosis; rather, they likely participate in the fine-tuning of presynaptic activity (Sun et al., 2006).

3.1.4 Rab3A The rab family small GTP-binding proteins are widely involved in vesicular trafficking. The active form of each rab protein (i.e., the GTP-form) is bound to the vesicles or the organelles via attached prenyl groups. In the synaptic vesicles, rab3A is the main rab protein and many rab3A-interacting proteins are characterized (Li and Chin, 2003; Su¨dhof, 2004). Although a number of studies, including analyses of knockout mice, indicate that rab3A and its activator, rab3A-GEF, are indispensable for transmitter release (Yamaguchi et al., 2002), the specific role of rab3A in exocytosis remains unknown.

3.1.5 Vacuolar-ATPase Vacuolar-ATPase (V-ATPase) is an H+-ATPase. The V-ATPase is composed of V0 (260 kDa; membraneinserted H+-channel) and V1 (600–650 kDa; ATPase) domains. Each domain of V-ATPase contains multiple subunits, one of which hydrolyzes one molecule of ATP, resulting in the uptake of two H+ into the vesicles (Inoue et al., 2005). This ATP-dependent H+ concentration gradient is used to take up and condense various neurotransmitters into the SVs by active transport.

3.1.6 Cysteine String Protein Cysteine string protein (CSP) is a member of DnaJ chaperone family. This protein acts as a chaperone for the Ca2+ channels in cooperation with other chaperones such as Hsc70 (Evans et al., 2003). Knockout mouse studies suggest that CSP plays an important role in presynaptic terminal functions (Chandra et al., 2005).

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3.2 Active Zone Proteins The active zone (AZ) is the site where the synaptic vesicles accumulate and where exocytosis frequently occurs. Thus, proteins in the AZ are expected to participate in vesicle accumulation and the stimulation of exocytosis (Schoch and Gundelfinger, 2006; > Figure 5‐3). Recent studies reveal that a large protein complex is formed in the AZ through interactions between the various AZ proteins.

. Figure 5‐3 Proteins localized in the AZ and their interactions. AZ proteins (yellow) interact with each other, adapter proteins (blue), synaptic membrane proteins (pink), and cytoskeletal proteins (brown)

3.2.1 Munc13 Munc13, a large protein with a single C1 phorbol ester-binding domain and three C2 domains, is a homolog of Unc-13 in C. elegans and acts at the vesicle priming step by interacting with nematode syntaxin Unc-64 (Richmond et al., 2001; Li and Chin, 2003). There are four isoforms of Munc13 named Munc13-1, ubMunc13-2, bMunc13-2, and Munc13-3. These proteins are key regulators of presynaptic short-term plasticity, a process where neurotransmitter release is dynamically adapted to conditions. The isoform ubMunc13-2 is ubiquitous, whereas the others are restricted to the brain. Both Munc13-1 and ubMunc13-2 bind calmodulin (CaM) in a Ca2+-dependent manner (Junge et al., 2004). The steady-state excitatory presynaptic current (EPSC) amplitude under conditions of vesicle depletion, representing the rate of rapidly releasable pool (RRP) refilling, was lower in mice expressing a W387R mutant of Munc13-1 or ubMunc13-2, which cannot bind CaM, than in wild-type mice. These lower steady-state amplitudes are rescued with Munc13-1 (W464R) and indicate impaired priming activity, which, in turn, alters short-term plasticity (Junge et al., 2004). In addition, the accelerated RRP refilling and the accompanying increase in RRP size is impaired by preventing CaM-Munc13 binding. It appears that CaM-Munc13 interactions are involved in the regulation of activity-dependent vesicle priming.

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3.2.2 Bassoon and Piccolo These two proteins are large and contain multiple domains. Both contain two zinc-finger domains and three coiled-coil domains, and piccolo has one PDZ domain and two C2 domains (Garner et al., 2000). These two proteins act as scaffolds for the formation of protein complexes by mediating multiple protein–protein interactions (see Schoch and Gundelfinger, 2006).

3.2.3 CAST/ELKs CAST (also known as ELK, ERC, and rab6-interacting protein) directly forms a large complex with RIM, bassoon, and piccolo and indirectly interacts with Munc13-1 via RIM1, all of which are AZ components (Takao-Rikitsu et al., 2004). In cultured mammalian neurons, this protein must localize in the AZ and is involved in synaptic activity. Thus, CAST is thought to be one of the key proteins mediating the organization of the AZ.

3.2.4 RIM RIM is a homolog of C. elegans unc-10 and interacts with rab3A (Wang et al., 1997). RIM is also a multidomain protein that binds AZ components such as Munc13s, CAST, bassoon, and piccolo (Schoch and Gundelfinger, 2006). Knockout mice show, however, that RIM is not an indispensable component of the AZ. RIM1a null mice-derived autapses have a reduced readily releasable pool (Calakos et al., 2004) and changes in the properties of evoked asynchronous release. Phosphorylation of RIM1a by A-kinase or by SAD kinase are shown to be important for modulation of exocytosis (Inoue et al., 2006).

3.3 Presynaptic Membrane Proteins There are several presynaptic membrane proteins other than t-SNAREs that participate in synaptic function and formation (> Figure 5‐3).

3.3.1 Neurexins The first neurexin was discovered as a Ca2+-dependent receptor for a-latrotoxin, a spider toxin that forces exocytosis from the synapse (Ushkaryov et al., 2004). The neurexin family of proteins is believed to be partly involved in the stabilization of immature excitatory synapses through an interaction with the postsynaptic protein neuroligin (Craig and Kang, 2007). Neurexin also interacts with the Ca2+ channel (Missler et al., 2003).

3.3.2 Ca2+ Channels Ca2+ channels are composed of five subunits (a1, a2, b, g, and d). The large a1 subunit serves as the conduit for Ca2+. The Ca2+ channels are also physiologically and pharmacologically classified into several groups, of which the neuronal forms include N, P/Q, and R. For example, the Cav2.1 a1 subunit corresponds to the P/Q-type channel and Cav2.2 to the N-type channel. Ca2+ channels are characterized according to their a1 subunits. Of the other subunits, the b-subunit is localized at the cytoplasmic face and is known to interact with signaling proteins (Evans and Zamponi, 2006). Ca2+ channels interact with H3 domains of the syntaxin-1A via the synprint sequence in their a1 subunits, and this mechanism is believed to link the Ca2+ channels to the fusion machinery (Evans and Zamponi, 2006). CaM is known to be bound to the a1 subunit at two different binding sites, probably to sense Ca2+ concentrations (Evans and Zamponi, 2006). Because the clustering of the Ca2+

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channels in the presynaptic terminals are critical for massive, synchronized, and rapid (ms) release, the mechanism of their accumulation is very important. Although the molecular mechanisms are not completely clarified, some possibilities include binding by RIM-binding protein and neurexin (Hibino et al., 2002).

4

The Proteins Involved in Ca2+-Dependent Vesicular Recycling

4.1 Synaptotagmins and Low-Affinity Ca2+ Sensors Low-affinity Ca2+ sensors mediate fusion between the lipid bilayers of SVs and the plasma membrane. Synaptotagmin family proteins, specifically those that bind phospholipids and contain two C2 domains, which bind Ca2+ with micromolar affinity, are thought to be such fusion-mediating Ca2+ sensors (Su¨dhof, 2004). Because v-SNARE cannot access inappropriate t-SNARE complexes formed in the plasma membrane, a high concentration of Ca2+ necessary to cause fusion, which may help prevent the formation of inappropriate complexes, or it may maintain the appropriate SNARE complex in an energetically more favorable state. In addition, rapid fusion occurs at site where Ca2+ channels are clustered (Wadel et al., 2007). Taken together, it is likely that the interaction between t-SNARE proteins and Ca2+ channel clusters (Catterall, 1998) before docking may regulate the above ‘‘competent state’’ of the SVs.

4.2 Ca2+-Dependent Vesicular Recycling Regulated exocytosis is separated into several steps: (1) SV recruitment beneath the plasma membrane, (2) tethering, (3) docking, (4) priming, and (5) vesicular fusion (> Figure 5‐4). These steps are characterized by electrophysiological studies, but the molecular complexes involved are not fully understood. Some steps are mediated by different concentrations of Ca2+. A tethering process occurs prior to formation of the SNARE complex (Li and Chin, 2003). This process shortens the distance between the vesicle and the target membrane. Tethering in presynaptic exocytosis is not sufficiently characterized, but it is well-understood for the transport of other vesicles in the Golgi apparatus (Gillingham and Munro, 2003; Lupashin and Sztul, 2005).

4.3 Submicromolar Ca2+ Concentrations Regulate Vesicular Recycling SV fusion requires 1–10 mM Ca2+ (Burgoyne and Morgan, 2003), but it is also required for endocytosis and the supply of vesicles for neurotransmission. The latter process requires submicromolar concentrations (0.3–0.4 mM) of Ca2+, which is slightly higher than the resting Ca2+ concentration (> Figure 5‐4; Burgoyne and Morgan, 2003). High-affinity Ca2+ sensors have not yet been identified. CaM has four Ca2+-binding sites (Jurado et al., 1999) and is not activated at a resting Ca2+ levels, but it is fully activated by 0.5 mM Ca2+. CaM binds Ca2+ cooperatively, that is, if one atom of Ca2+ is bound, the affinity of the other Ca2+-binding sites, even the low-affinity sites, increases (Burgoyne and Clague, 2003). Putative low-affinity Ca2+ sensors for vesicular fusion require the high off rate Ca2+ dissociation within 1 ms after Ca2+ release (Burgoyne and Clague, 2003), and it does not act as the Ca2+ sensor mediating rapid fusion. Instead, CaM can mediate submicromolar Ca2+-dependent intracellular vesicular trafficking (Burgoyne and Clague, 2003). Sakaba and Neher (2001) found that the transfer of vesicles from the reserve pool (RP) to the RRP is inhibited by a CaM inhibitor in neurons. Their results show that CaM is not directly involved in the Ca2+-triggered release process (i.e., SV fusion) but instead is involved in short-term plasticity. We discuss the roles of CaM-binding proteins in SV recycling later.

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. Figure 5‐4 Vesicle trafficking in the presynaptic terminal. In the presynaptic terminal, SVs are physiologically classified into two groups: (a) the RRP, which are vesicles used for rapid release and response to the initial stimulation; and (b) the RP, which are vesicles resupplying the RRP after frequent stimulation. The RRP vesicles are believed to be ‘‘docked’’ (immobilized on the plasma membrane probably through the SNARE complex) and/or ‘‘primed’’ (more readily fused due to biochemical modulation of the SNARE complex) for release at submicromolar concentrations of intracellular Ca2+ (Stojilkovic, 2005). A higher concentration of Ca2+ (>1 mM) is thought to be required for vesicular fusion. After endocytosis, the vesicles are recycled and are part of the RP. Currently, the molecular mechanisms of the transition from the RP to RRP remain unknown. It is thought that tethering occurs prior to the docking step; however, its role in the synaptic terminal is under investigation. We propose that the interaction of myosin-V and syntaxin-1A (Watanabe et al., 2005) is one of the potential candidates explaining this step (see > Figure 5‐6)

CaM is not the only Ca2+-binding protein involved in exocytosis. Neuronal calcium sensor (NCS)-1/frequenin is a member of NCS family and has an EF-hand Ca2+-binding motif with higher affinity for Ca2+ than CaM. This protein has a submicromolar Kd for Ca2+ (0.3 mM) in vitro and participates in exocytosis via the regulation of Ca2+ channels (Tsujimoto et al., 2002). Although each of the NCS family members is much less abundant than CaM in neurons, they may contribute to specific presynaptic events.

4.4 CaM-Dependent Regulation of Exocytosis Through Modulation of Membrane Trafficking Proteins We reasoned that if submicromolar concentrations of Ca2+ regulate vesicular recycling in neurons, there should be protein–protein interactions responding to these same concentrations of Ca2+ that regulate the SNARE complex. We focused on syntaxin-1A as a possible target for regulation by submicromolar Ca2+ and discovered two Ca2+-dependent syntaxin-binding proteins, CaMKII and myosin-V (Ohyama et al., 2002; Nomura et al., 2003; Watanabe et al., 2005).

4.4.1 CaMKII CaMKII functions as a complex of 8–12 monomers and is activated by Ca2+/CaM at a concentration of Ca2+ less than 1 mM through the autophosphorylation of one monomer (Hudmon and Schulman, 2002;

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Lisman et al., 2002). The decrease of Ca2+ reduces the probability of CaMKII autophosphorylation (Hudmon and Schulman, 2002) and activates phosphatases that dephosphorylates CaMKII and inactivates it (Hudmon and Schulman, 2002; Lisman et al., 2002). In the presynaptic terminal, CaMKII is one of the major SV proteins. We demonstrated that only autophosphorylated CaMKII binds to syntaxin-1A in the presence of submicromolar Ca2+. Both decreased Ca2+ and the dephosphorylation of CaMKII induced the reversible release of CaMKII from syntaxin-1A. We found that the binding of CaMKII occurs in the linker domain of syntaxin-1A (Ohyama et al., 2002) and currently, CaMKII is the only protein known to bind this site (Ohyama et al., 2002; Nomura et al., 2003). Because the CaMKII complex is as large as 500 kDa and because the linker domain regulates the conformation of syntaxin, it must fix bound syntaxin in the open form. We biochemically confirmed that the CaMKII-bound syntaxin-1A is in the open form and that this complex recruits SNAP-25 and synaptotagmin to form the SNARE complex. Thus, the CaMKII–syntaxin complex appears to take on the open form, which promotes formation of the SNARE complex, a process essential for exocytosis (> Figure 5‐5). . Figure 5‐5 Significance of the CaMKII-syntaxin-1A interaction. Autophosphorylated CaMKII (P-CaMKII), activated by the binding of Ca2+/CaM (the large S-shape), can bind the linker domain in presence of 1 mM Ca2+(Ohyama et al., 2002). This complex can then recruit SNAP-25 and synaptotagmin (Stg), allowing it to smoothly shift to the docking complex. Because the linker domain governs the interconversion between the open and the closed forms, binding of CaMKII to this domain is likely to fix the syntaxin molecule (Syx) in the open form, facilitating formation of the SNARE complex. Munc-18 is a protein that fixes bound syntaxin in the closed form

Chronic treatment with antidepressants has been shown to decrease glutamate release and decrease the amount of syntaxin 1A-phosphorylated CaMKII complex, while increasing the amount of syntaxin 1A–Munc-18 complex, suggesting that antidepressants control transmitter release by regulating CaMKIIdependent SNARE protein interactions (Bonanno et al., 2005).

4.4.2 Myosin-V Myosin-V, an unconventional myosin, acts as a molecular motor for the local intracellular transport of vesicles and organelles along F-actin (Reck-Peterson et al., 2000; Vale, 2003). In cortical neurons, myosin-V is localized around the SVs (Tilelli et al., 2003) and appears to regulate vesicle recycling. We found that myosin-V binds to syntaxin-1A in the presence of submicromolar Ca2+ (Watanabe et al., 2005). The most important aspect of this interaction is that this binding occurs in the neck domain of myosin-V, which is where CaM is attached as light chains. Myosin-V releases one-sixth of CaM from the

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neck domain in presence of submicromolar Ca2+ (Cameron et al., 1998). This Ca2+-dependent CaM release is linked to binding of syntaxin-1A to the neck region (> Figure 5‐6a). Inhibition of this interaction decreased the probability of exocytosis in chromaffin cells, confirming that this interaction is involved in exocytosis (Watanabe et al., 2005) and in RP- but not RRP-dependent exocytosis (Watanabe et al., 2005).

. Figure 5‐6 A putative model for vesicular transport regulated by myosin-V. (a) Model of the complex between myosin-V and syntaxin-1A (Watanabe et al., 2005). Each monomer of myosin-V is composed of a head region, a long neck domain containing six tandem IQ-motifs, and a tail region. The light chains consist mainly of CaM bound to the IQ-motifs in the neck domain. The tail interacts with a receptor protein on the conveyed cargoes (i.e., organelles and vesicles). The neck region of myosin-Va senses the elevation in Ca2+ via its neck region and exchanges one CaM molecule for one syntaxin-1A molecule. (b) Our model of myosin-V-dependent vesicular recycling (Watanabe et al., 2005). Myosin-V, which mediates F-actin-dependent conveyance to the plasma membrane, is on the surface of the vesicle and binds to syntaxin-1A when the intracellular Ca2+ concentration increases to 0.3–0.4 mM. After the complex between myosin-V and syntaxin-1A tethers the vesicle to the membrane, it recruits other SNARE proteins to form the SNARE complex. A second larger elevation in [Ca2+] induces vesicular fusion (Rettig and Neher, 2002) and stimulates the exchange of myosin-V for NSF/SNAP in the SNARE complex. Myosin-VI, a reverse motor, mediates endocytosis (Hasson, 2003), whereas myosin-V, an orthotropic motor, probably participates in exocytosis

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Therefore, this interaction is concluded to participate in the translocation of the secretory vesicles used for exocytosis (cf. Desnos et al., 2007). Myosin-V on the vesicles is large, flexible, and contains coiled-coil regions that could mediate tethering (Gillingham and Munro, 2003; Lupashin and Sztul, 2005). Because myosin-V interacts with syntaxin-1A through the neck region, during conveyance of the vesicle, myosin-V can interact with syntaxin-1A in the plasma membrane, bringing the vesicle close and therefore tethering it to the plasma membrane (> Figure 5‐6b). This hypothesis is also supported by a recent report showing that chromaffin granules directly interact with the membrane via an interaction between myosin-V and TIRFM. Although dilute mutant mice, which lacks in myosin-Va expression, do not show obvious changes in exocytosis (Schnell and Nicoll, 2001), two recent reports indicate a reduction of exocytotic activity in retinal and hippocampal neurons of dilute mice (Trinchese et al., 2003; Libby et al., 2004).

4.4.3 VAMP/Synaptobrevin and CaM VAMP-2 binds to CaM in a Ca2+-dependent manner, which can regulate the acidic phospholipid-binding activity of VAMP (Quetglas et al., 2002). CaM binding also prevents VAMP from forming the SNARE complex (Quetglas et al., 2002). The CaM-binding site of VAMP (77–90) lowers the exocytotic frequency (Quetglas et al., 2002). These results suggest that CaM modulates the function of VAMP by regulating vesicle fusion (de Haro et al., 2004).

5

Conclusions

In this review, we discuss how proteins localized in the presynaptic area are effectively arranged. However, the roles of a number of the presynaptic proteins remain unclear. Because the complexity of presynaptic functions probably arises from molecular redundancy and the variety of protein isoforms, more precise molecular interaction maps between these proteins are needed. In addition, the presynaptic events that have been electrophysiologically confirmed to occur at given Ca2+ concentrations remain to be explained according to the specific Ca2+-dependent protein interactions. Recent studies are beginning to elucidate the molecular aspects of how CaM contributes to submicromolar Ca2+-dependent events. Because SV recycling is extremely rapid and because more than ten species of proteins are involved, it is difficult to determine how the molecular machinery functions. Further studies using bioimaging and gene targeting technologies should help in this regard.

Acknowledgments This work was in part supported by grants from the Ministry of Education, Culture, Science, Sports, and Technology of Japan (#16044216; #17023019) and by grants to M.I. from the Project Promoting Program and the Strategic Research Program of the Niigata University. This article is dedicated to the late Yoshiaki Komiya (Emeritus Professor, Gunma University) for his encouragement when M.I. started in this field of research.

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Schnell E, Nicoll RA. 2001. Hippocampal synaptic transmission and plasticity are preserved in myosin Va mutant mice. J Neurophysiol 85: 1498-1501. Schoch S, Gundelfinger ED. 2006. Molecular organization of the presynaptic active zone. Cell Tissue Res 326: 379-391. Shen J, Tareste DC, Paumet F, Rothman JE, Media. TJ2007. Selective activation of cognate SNAREpins by Sec1/Munc18 proteins. Cell 128: 183-195. Sieber JJ, Willig KI, Kutzner C, Gerding-Reimers C, Harke B, et al. 2007. Anatomy and dynamics of a suparmolecular membrane protein cluster. Science 317: 1072-1076. Stojilkovic SS. 2005. Ca2+-regulated exocytosis and SNARE function. Trends Endocrinol Metabol 16: 81-83. Su¨dhof TC. 2002. Synaptotagmins: Why so many? J Biol Chem 277: 7629-7632. Su¨dhof TC. 2004. The synaptic vesicle cycle. Annu Rev Neurosci 27: 109-147. Sun J, Bronk P, Liu X, Han W, Su¨dhof TC. 2006. Synapsins regulate use-dependent synaptic plasticity in the calyx of Held by a Ca2+/calmodulin-dependent pathway. Proc Natl Acad Sci USA 103: 2880-2885. Takamori S, Holt M, Stenius K, Lemke EA, Gronborg M, et al. 2006. Molecular anatomy of a trafficking organelle. Cell 127: 831-846. Takao-Rikitsu E, Mochida S, Inoue E, Deguchi-Tawarada M, Inoue M, et al. 2004. Physical and functional interaction of the active zone proteins, CAST, RIM1, and bassoon, in neurotransmitter release. J Cell Biol 164: 301-311. Tang J, Maximov A, Shin OH, Dai H, Rizo J, et al. 2006. A complexin/synaptotagmin 1 switch controls fast synaptic vesicle exocytosis. Cell 126: 1175-1187. Tilelli CQ, Martins AR, Larson RE, Garcia-Cairasco N. 2003. Immunohistochemical localization of myosin Va in the adult rat brain. Neuroscience 121: 573-586. Tokumaru H, Umayahara K, Pellegrini LL, Ishizuka T, Saisu H, et al. 2001. SNARE complex oligomerization by synaphin/complexin is essential for synaptic vesicle exocytosis. Cell 104(3): 421-432. Toonen RF, Kochubey O, de Wit H, Gulyas-Kovacs A, Konijnenburg B, et al. 2006. Dissecting docking and tethering of secretory vesicles at the target membrane. EMBO J 25: 3725-3737. Trinchese F, Rao M, Peterhoff C, Kumar A, Liu S, et al. 2003. Myosin Va is required for neurotransmitter release during basal synaptic transmission and synaptic plasticity. Mol Biol Cell 14: 179a. Tsujimoto T, Jeromin A, Saitoh N, Roder JC, Takahashi T. 2002. Neuronal calcium sensor 1 and activity-dependent facilitation of P/Q-type calcium currents at presynaptic nerve terminals. Science 295: 2276-2279.

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Ushkaryov YA, Volynski KE, Ashton AC. 2004. The multiple actions of black widow spider toxins and their selective use in neurosecretion studies. Toxicon 43: 527-542. Vale RD. 2003. The molecular motor toolbox for intracellular transport. Cell 112: 467-480. Verhage M, Maia AS, Plomp JJ, Brussaard AB, Heeroma JH, et al. 2000. Synaptic assembly of the brain in the absence of neurotransmitter secretion. Science 287: 864-867. Wadel K, Neher E, Sakaba T. 2007. The coupling between synaptic vesicles and Ca2+ channels determines fast neurotransmitter release. Neuron 53: 563-575.

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6

Synaptic Plasticity in the Cerebellum

T. Tabata . M. Kano

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64

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Synaptic Organization of the Cerebellum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65

3 3.1 3.2 3.3

Plasticity of parallel fiber (PF)-Purkinje cell (PC) Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Paired-Pulse Facilitation (PPF) at PF-PC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Short-Term Potentiation (STP) at PF-PC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Short-Term Depression (STD) at PF-PC Synapses: Endocannabinoid-Mediated Retrograde Depression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Long-Term Potentiation (LTP) at PF-PC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Long-term depression (LTD) at PF-PC Synapses: Cerebellar LTD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5.1 What is Cerebellar LTD? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5.2 Molecular Mechanisms Underlying Cerebellar LTD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

66 66 67

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Plasticity of CF-PC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Paired-Pulse Depression (PPD) at CF-PC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Depolarization-induced suppression of excitation (DSE) at CF-PC Synapses . . . . . . . . . . . . . . . . . . . LTD at CF-PC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

74 74 74 75

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Plasticity of IN-PC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . STP at interneuron (IN)-PC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . STD at IN-PC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . LTP at IN-PC Synapses: Rebound Potentiation (RP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

75 75 75 75

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Plasticity of PF-IN Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 STD at PF-SC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 LTP/LTD at PF-SC Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

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LTP at mossy fiber (MF)-deep cerebellar nuclear neuron (DCNN) Synapses . . . . . . . . . . . . . . . . . 77

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LTD at PC-DCNN Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77

9 9.1 9.2 9.3 9.4 9.5

Physiological Significance of Cerebellar Synaptic Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Modern Behavioral Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Motor Coordination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Delayed Eye-blink Conditioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Adaptation of Vestibulo-Ocular Reflex (VOR) and Optokinetic Reflex (OKR) . . . . . . . . . . . . . . . . . Fear Conditioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Conclusive Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83

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Synaptic plasticity in the cerebellum

Abstract: Certain patterns of neuronal activities induce changes in the efficacy of synaptic transmission. These phenomena are called synaptic plasticity, and assumed as a physiological basis for learning and memory. The cerebellum provides a powerful experimental paradigm for studying synaptic plasticity. The synaptic circuits of the cerebellum consist of the cerebellar cortex and deep cerebellar nuclei. The cerebellar cortex is a stack of stereotyped circuitry containing only a few types of neurons. Besides, the pattern of projections between the cerebellar cortex, deep cerebellar nuclei, and related brain regions is relatively simple. This allows an analysis of synapses between definitely identified neuronal types. The cerebellum is shown to be important for some sorts of motor learning. Thus, one can pursue the physiological significance of cerebellar synaptic plasticity at the behavioral level, using motor learning tasks. To date, various forms of synaptic plasticity have been demonstrated at distinct cerebellar synapses. In this chapter, we review some representative forms of cerebellar synaptic plasticity and discuss their possible mechanisms and physiological significance. List of Abbreviations: AMPAR, (RS)-a-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid-type ionotropic glutamate receptor; BC, basket cell; CB1R, CB1 cannabinoid receptor; CF, climbing fiber; CRF, corticotrophin-releasing factor; DAG, diacylglycerol; DCN, deep cerebellar nuclei; DCNN, deep cerebellar nuclear neuron; DSE, depolarization-induced suppression of excitation; DSI, depolarization-induced suppression of inhibition; EPSC, excitatory postsynaptic current; EPSPs, excitatory postsynaptic potentials; ERK1/2, extracellular signal-regulated kinase; GC, granule cell; GRIP, glutamate receptor-interacting protein; IN, interneuron; IP3, inositol trisphosphate; IP3R, IP3 receptor; IPSPs, inhibitory postsynaptic potentials; LTD, long-term depression; LTP, long-term potentiation; MEK1/2, mitogen-activated protein kinase; MF, mossy fiber; mGluR1, type-1 metabotropic glutamate receptor; NMDAR, N-methyl-D-aspartate-type ionotropic glutamate receptor; OKR, optokinetic reflex; PC, Purkinje cell; PF, parallel fiber; PKA, protein kinase A; PKC, protein kinase C; PPD, paired-pulse depression; PPF, paired-pulse facilitation; RP, rebound potentiation; SC, stellate cell; sGC, soluble guanylyl cyclase; STD, short-term depression; STP, short-term potentiation; VGCCs, voltage-gated Ca2+ channels; VOR, vestibulo-ocular reflex

1

Introduction

It is widely believed that synaptic plasticity is a physiological basis for learning and memory. Synaptic plasticity is a change in the efficacy of synaptic transmission induced by a certain pattern of neuronal activity. Synaptic plasticity could modify information flow in the brain, and thereby consolidate memory traces. The cerebellum, a brain structure profoundly involved in motor learning (see > Section 2), has long been a ‘‘hot spot’’ for studying synaptic plasticity. By the end of the 1960s, the detailed diagram of the cerebellar circuit was depicted by the pioneers including John Eccles (see Kano et al., 2007 for the review of the history of functional cerebellar research). Around 1970, David Marr and James Albus independently proposed network theories that temporally correlated activities of two classes of excitatory input fibers to induce long-term synaptic plasticity in the cerebellar cortex (see > Section 2 for the anatomical terms). Assuming such a synaptic plasticity, Masao Ito hypothesized that the cerebellar flocculus (an evolutionally old part of the cerebellar cortex) adaptively controls vestibulo-ocular reflex (see > Section 9.4). In the late 1970s, there was increasing circumstantial evidence for the validity of the plasticity assumption. In 1982, Ito and coworkers eventually demonstrated a form of synaptic plasticity in the cerebellar cortex in vivo and termed it cerebellar long-term depression (LTD) (see > Section 3.5.1). In early studies on cerebellar LTD, unit or field neuronal activities were recorded in vivo, using relatively large mammals such as rabbits. In 1987, Masaki Sakurai succeeded in measuring cerebellar LTD in situ, using cerebellar slices. In 1990, Arthur Konnerth and coworkers established a technique for patch-clamp recordings from neurons in cerebellar slices. Around the same time, Tomoo Hirano succeeded in inducing cerebellar LTD-like phenomena in vitro in cultured cerebellar neurons. The use of these in situ and in vitro preparations offers great technical advantages. In these preparations, one can visually identify various types of cerbellar neurons under microscopy, and make recordings from neurons of interest without difficulty.

Synaptic plasticity in the cerebellum

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This led not only to a detailed characterization of cerebellar LTD, but also to the discovery of many other forms of cerebellar synaptic plasticity. Moreover, one can readily deliver pharmacological agents to cerebellar neurons by perfusing the slice or cultured neurons with drug-containing saline. Cerebellar slices and cultured cerebellar neurons can be prepared from relatively small animals including genetically modified mice. These enable an analysis of cerebellar synaptic plasticity at the molecular level. On the other hand, the physiological significance of cerebellar synaptic plasticity has been pursued by examining animals’ performance in motor and other behavioral tasks. Classical behavioral studies employing cerebellar lesions indicated the importance of the cerebellum for motor control and learning. Recent behavioral studies attempt to elucidate the precise roles of cerebellar synaptic plasticity, employing genetic manipulation of plasticity-related molecules. In this chapter, we review some representative forms of cerebellar synaptic plasticity described to date, and discuss their possible mechanisms and physiological significance.

2

Synaptic Organization of the Cerebellum

The synaptic circuits of the cerebellum consist of the cerebellar cortex and deep cerebellar nuclei (DCN) (> Figure 6-1a,b). As their names imply, the cerebellar cortex is a neuron-rich structure covering the outer surface of the cerebellum and the DCN are neuron-rich structures deep inside the cerebellum. The cerebellar cortex contains a stack of stereotyped circuitry consisting of only a few types of neurons (> Figure 6-1b,c). The DNC are an archipelago of small nuclei: the fastigial nucleus, interposed nuclei, and dentate nucleus. The cerebellar cortex receives two classes of excitatory inputs: mossy fibers (MFs) and climbing fibers (CFs) (> Figure 6-1c) (Ito, 2006). MFs originate in the spinal cord and brainstem. CFs are the axons of inferior olive neurons. MFs convey sensory information from various body parts and motor commands from the upper centers. These signals are transferred to granule cells (GCs) via glutamatergic (excitatory) synapses (> Figure 6-1b,c). The axons of GCs [parallel fibers (PFs)] give off their branches running laterally in the molecular layer of the cerebellar cortex. Purkinje cells (PCs) spread huge, flat dendritic arbors along the parasagittal planes in the molecular layer. Each PC receives excitatory synaptic inputs from more than 100,000 PFs. PCs integrate sensory information and motor commands through summation of PF inputs. PCs are the sole output neurons of the cerebellar cortex and send the integrated signals to DCN neurons (DCNNs) and vestibular nuclear neurons via GABAergic (inhibitory) synapses. PFs also form glutamatergic synapses on interneurons (INs) in the cerebellar cortex. INs include Golgi cell, stellate cell (SC), and basket cell (BC). Golgi cells form inhibitory synapses on the synaptic terminals of MFs. The PF-Golgi cell-MF loop circuitry imposes feedback inhibition, which attenuates glutamate release by the PFs. SCs and BCs form inhibitory synapses on PCs. A single set of stimulation of a group of PFs first evokes excitatory postsynaptic potentials (EPSPs) and then, with a short delay, evokes inhibitory postsynaptic potentials (IPSPs) in the PCs. The latter are mediated by the PF-SC/BC-PC circuitry and may truncate the preceding PF-PC EPSPs (feedforward inhibition). The feedforward inhibition limits the time-window for PF-PC EPSPs to contribute to the generation of action potentials in PCs. Each PC is innervated by a single CF. CFs convey error signals reporting a difference between intended and executed movements. CF-PC EPSPs serve as a trigger for some forms of synaptic plasticity. DCNNs integrate excitatory synaptic inputs from MFs and inhibitory synaptic inputs from PCs, and send the integrated signals directly to the red nucleus and indirectly to the primary motor cortex and premotor cortex via the thalamus. The cerebellum influences the regulatory actions of these nuclei and cortices on the descending motor system, and thereby contributes to motor coordination and motor accuracy. Plasticity at synapses upstream of PCs changes the transferring efficacy of sensory information and motor commands to PCs. Plasticity of synapses on PCs and DCNNs changes the manner of signal integration in these neurons. Cerebellar synaptic plasticity may thereby further improve motor coordination and motor accuracy, and contribute to the acquisition of new motor skills.

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. Figure 6-1 Anatomical and functional structure of the cerebellum. (a) Locations of the cerebellum and its related regions shown in a midline-sectioned human brain. Adapted from (Purves, 2004). (b) 3D structure of single folia of the cerebellar cortex. Adapted from (Purves, 2004). (c) Basic circuitry of the cerebellum. BC, basket cell; CC, cerebellar cortex; CF, climbing fiber; CN, (deep) cerebellar nucleus; GL, glomeruli; GO, Golgi cell; GR: granule cell; IO, inferior olive; LC, Lugaro cell; MF, mossy fiber; N-C, nucleo-cortical mossy fiber projection; N-O, nucleoolivary inhibitory projection; PC, Purkinje cell; PCN, precerebellar neuron; PF, parallel fiber; pRN, parvicellular red nucleus; R-O, rubro-olivary excitatory projection; SC, stellate cell; SR, serotonergic fiber; UB, unipolar brush cell; VN, vestibular nucleus. Adapted from (Ito, 2006)

3

Plasticity of parallel fiber (PF)-Purkinje cell (PC) Synapses

3.1 Paired-Pulse Facilitation (PPF) at PF-PC Synapses When a pair of electrical pulses is given to PFs with a short interval (10–500 ms), the amplitude of PF-PC excitatory postsynaptic current (EPSC) evoked by the second pulse exceeds that of PF-PC EPSC evoked by the first pulse (> Figure 6-2a) (Atluri and Regehr, 1996). Such homosynaptically induced, short-lived potentiation is called PPF.

Synaptic plasticity in the cerebellum

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. Figure 6-2 PPF and PPD at the cerebellar synapses. (a) PPF of PF-PC EPSC. Plot: The ratio of amplitudes of PF-PC EPSCs evoked by a pair of electrical pulses to the PFs as a function of inter-pulse interval. Inset: sample EPSCs evoked with an interval of 50 ms. Adapted from (Atluri and Regehr, 1996). (b) PPD of CF–PC EPSC. Sample EPSCs evoked with an interval of 40 ms. Adapted from (Hashimoto and Kano, 1998)

In PFs, an action potential(s) evoked by an electrical pulse opens voltage-gated Ca2+ channels (VGCCs). Extracellular Ca2+ enters through the VGCCs and may activate a signaling molecule(s) that exerts a sustained enhancing effect on neurotransmitter release (Atluri and Regehr, 1996). Candidates for such a molecule include calmodulin and synaptotagmin III. Because of its presynaptic origin, PPF of PF-PC EPSC can be utilized as a measure to monitor the modulation of the presynaptic release machinery.

3.2 Short-Term Potentiation (STP) at PF-PC Synapses Periodic burst stimuli (e.g., 90 sets of bursts at one Hz, each burst consists of 5 pulses at 50 Hz) to PFs induce short-term potentiation (STP) of PF-PC EPSP, lasting for 10–20 min (Goto et al., 2006). This form of STP may be due to the modulation of the presynaptic mechanisms (Goto et al., 2006). GCs discharge at a rate of 10–50 Hz at rest and 50–100 Hz in response to sensory stimuli in vivo (Ito, 2001; Chadderton et al., 2004). Thus, this form of STP may occur under physiological conditions.

3.3 Short-Term Depression (STD) at PF-PC Synapses: Endocannabinoid-Mediated Retrograde Depression In the cerebellum and hippocampus, a prolonged depolarizing stimulus (1 to a few s) to some postsynaptic neurons induces short-term depression (STD) (10–20 s) of excitatory or inhibitory neurotransmitter release by the presynaptic neurons (depolarization-induced suppression of excitation or inhibition; DSE or DSI, respectively) (see Chevaleyre et al., 2006; Hashimotodani et al., 2007 for review). DSE occurs at PF-PC, PF-SC, PF-BC, and CF-PC synapses (Kreitzer and Regehr, 2001; Maejima et al., 2005; Beierlein and Regehr, 2006). In the depolarized neurons, extracellular Ca2+ enters through VGCCs and then triggers the production of endocannabinoids such as 2-arachidonoylglycerol (2-AG) (> Figure 6-3a). Endocannabionids are naturally occurring analogs of the main component of marijuana. Endocannabinoids diffuse through the plasma membrane and bind retrogradely to presynaptic CB1 cannabinoid receptor (CB1R). CB1R in turn attenuates Ca2+ entry through VGCCs via Gi/o protein, and this results in a decrease of neurotransmitter release. The mechanisms underlying DSE also mediate homosynaptically induced STD of PF-PC EPSP (> Figure 6-3b). Upon high-frequency stimuli (e.g., 10 pulses at 100 Hz) are given to PFs innervating a PC whose membrane potential is monitored under current clamp (i.e., the PC can freely discharge), EPSPs

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. Figure 6-3 STD at PF-PC synapses. (a) DSE is mediated by endocannabinoid-dependent retrograde signaling. (b, c) Highfrequency PF stimulation (10 pulses at 100 Hz) induces STD of PF-PC EPSP (c) through mechanisms overlapping that of DSE (b). Upper traces in C: Sample EPSPs recorded before, 2 s after, and 30 s after the high-frequency stimuli are superimposed. Lower trace in C: A sample voltage response of a PC to the high-frequency stimuli. Plots in C: the amplitude of EPSP in the absence and presence of a CB1R antagonist as a function of time after the high-frequency stimuli. Panels A and B adapted from (Ohno-Shosaku et al., 2005). Panel C adapted from (Maejima et al., 2005)

are depressed for 10–20 s in the PC (Maejima et al., 2005) (> Figure 6-3c). Glutamate released by the PFs binds to type-1 metabotropic glutamate receptor (mGluR1, a G protein-coupled receptor) in the PC (Maejima et al., 2005) (> Figure 6-3b). High-frequency PF stimulation evokes a strong depolarization with Ca2+ spikes in the dendrites of the PC (Rancz and Hausser, 2006). Gq/11 protein activated by mGluR1, together with Ca2+ entering from the extracellular side, activates phospholipase C (PLC) b4 (Maejima et al., 2005). PLCb4 in turn produces diacylglycerol (DAG), which is further converted into 2-AG, the retrograde messenger triggering the modulation of the presynaptic release machinery. Endocannabinoid-dependent STD may serve as a negative feedback that prevents over-excitation of PCs by PF inputs.

3.4 Long-Term Potentiation (LTP) at PF-PC Synapses PF stimulation induces two distinct forms of long-term potentiation (LTP) of PF-PC EPSC, depending on the stimulation frequency. A prolonged train of low-frequency stimuli (e.g., 1 Hz for 300 s) induces LTP lasting over 50 min by modulating the postsynaptic mechanisms (Lev-Ram et al., 2002) (> Figure 6-4a,b). This form of LTP depends on NO produced by the PFs, but not cAMP, cGMP, or protein kinase C

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. Figure 6-4 LTP at PF-PC synapses. (a) Two distinct forms of LTP of PF-PC EPSC induced by low- and moderately highfrequency PF stimulation (300 pulses at 1 Hz and 120 pulses at 4 Hz, respectively). Plot: the amplitude of EPSC as a function of time after the LTP-inducing stimuli. Traces: sample EPSCs recorded before and after the LTPinducing stimuli are superimposed. Adapted from (Lev-Ram et al., 2002). (b, c) Different mechanisms underlying the distinct forms of LTP. See the text for further explanation. (d) LTP induced by low-frequency PF stimulation (1 Hz for 5 min, ‘‘PF’’) and LTD induced by conjunctive PF and CF stimulation (1 Hz for 5 min, ‘‘PF + PC’’) reverse each other. Adapted from (Coesmans et al., 2004)

(PKC) (Lev-Ram et al., 2002). NO may enhance the N-ethylmaleimide-sensitive factor (NSF, a vesicular trafficking-related protein)-dependent postsynaptic expression of (RS)-a-Amino-3-hydroxy-5-methyl-4isoxazolepropionic acid-type ionotropic glutamate receptor (AMPAR) (see > Section 3.5.2) (Kakegawa and Yuzaki, 2005). On the other hand, moderately high-frequency stimuli (100–120 pulses at 4–10 Hz) induce LTP lasting over 50 min by modulating the presynaptic mechanisms (Salin et al., 1996; Lev-Ram et al., 2002; Qiu and Knopfel, 2007) (> Figure 6-4a,c). This form of LTP may require an increase of cAMP, but not NO (Salin et al., 1996; Lev-Ram et al., 2002). Ca2+/calmodulin-sensitve adenylyl cyclase I may produce cAMP in response to Ca2+ entering through VGCCs (Salin et al., 1996). cAMP activates protein kinase A (PKA), which may in turn enhance glutamate release by phosphorylating RIM1a, a protein regulating synaptic vesicle (Castillo et al., 2002). N-methyl-D-aspartate-type ionotropic glutamate receptor (NMDAR) activation in any cellular element(s) other than PFs may also be required for this form of LTP (Qiu and Knopfel, 2007).

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Low-frequency stimulus-induced LTP can reverse LTD at PF-PC synapses (cerebellar LTD, > Section 3.5.1) (Lev-Ram et al., 2003; Coesmans et al., 2004) (> Figure 6-4d). Thus, this form of LTP might contribute to erasing the memory of LTD-dependent motor learning.

3.5 Long-term depression (LTD) at PF-PC Synapses: Cerebellar LTD 3.5.1 What is Cerebellar LTD? Upon a prolonged train of synchronized stimuli (100–600 sets at 1–4 Hz) to a certain group of PFs and a CF innervating the same PC, EPSPs/EPSCs mediated by these PFs are depressed over a few hours (Ito, 2002) (> Figure 6-5a,b). This form of LTD was the first form of synaptic plasticity discovered in the cerebellum and termed cerebellar LTD. Cerebellar LTD is input-specific (> Figure 6-5a, right); cerebellar LTD modifies the weighing of inputs from specific PFs and the manner of integration of sensory information and motor commands in the PC. Cerebellar LTD is triggered by CF inputs that convey motor error signals. For these features, cerebellar LTD is considered as a principal physiological basis for cerebellar motor learning.

3.5.2 Molecular Mechanisms Underlying Cerebellar LTD The mechanisms underlying cerebellar LTD have been analyzed extensively at the molecular level (> Figure 6-6) (see Ito, 2002 for review). PFs form synapses on the dendritic spines of PCs. Glutamate released by PFs binds to AMPAR and mGluR1 that localize to the tips and annuli of PC dendritic spines, respectively. AMPAR is a cation-selective receptor channel that carries a major part of PF-PC EPSPs/EPSCs (> Figure 6-5). Its down-regulation is the primary cause of cerebellar LTD (see the following section). mGluR1 is a G protein-coupled receptor whose signaling is essential for inducing cerebellar LTD. Pharmacological inhibition or genetic knock-out of mGluR1 (mGluR1-KO mice) abolishes cerebellar LTD (Aiba et al., 1994; Shigemoto et al., 1994). PCspecific genetic rescue of mGluR1 in the mGluR1-KO mice (mGluR1-resuce mouse) restores cerebellar LTD (Ichise et al., 2000). In PC dendritic spines, mGluR1 colocalizes with B-type g-aminobutyric acid receptor (GABABR), a Gi/o protein-coupled receptor (Kulik et al., 2002). These receptors appear to form complexes (Tabata et al., 2004) (> Figure 6-7). GABABR enhances mGluR1 signaling in a Gi/o protein-independent manner in response to cerebrospinal fluid levels of GABA (a few tens of nanomolars) and/or Ca2+ (a few mM) (Tabata et al., 2004), and in a Gi/o protein-dependent manner in response to few mM GABA spilt over from neighboring inhibitory synapses (Hirono et al., 2001). These enhancements may increase the inducibility of cerebellar LTD (Tabata et al., 2004) (Kamikubo et al., a paper presented at the annual meeting of the Society for Neuroscience, 2007). Adenosine A1 receptor (A1R), another Gi/o proteincoupled receptor inhibits mGluR1 signaling in PCs in response to cerebrospinal levels (40–400 nM) of adenosine without the aid of Gi/o protein (Tabata et al., 2007). Upon ligand binding, mGluR1 activates Gq/11 protein (> Figure 6-6). Gq/11 protein in turn activates PLC. PLC cleaves phosphatidylinositol bisphosphate, a plasma membrane component, into DAG and inositol trisphosphate (IP3). IP3 opens IP3 receptor (IP3R), a ligand-gated Ca2+ channel in the endoplasmic reticulum (ER). Ca2+ stored in the ER effluxes to the cytoplasm through the IP3R. This intracellular Ca2+ mobilization is required for cerebellar LTD. Genetic knock-out of the dominant subtype of IP3R in PCs abolishes cerebellar LTD (Inoue et al., 1998). PFs release not only glutamate, but also NO in an activity-dependent manner (> Figure 6-6) (see Shibuki, 1993; Crepel, 1998 for review). NO diffuses through the plasma membrane and activates soluble guanylyl cyclase (sGC) in PCs. sGC produces cGMP, which in turn activates protein kinase G (PKG). The depletion of NO or inhibition of sGC abolishes cerebellar LTD, indicating that these molecules are required for cerebellar LTD.

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. Figure 6-5 Cerebellar LTD (LTD at PF-PC synapses). (a) A PC is innervated by numerous PFs and a single CF (left). Repetitive, temporally correlated activities of a certain group of the PFs (depicted as a single fiber for simplicity) and the CF (middle) induces LTD of PF-PC EPSP mediated by this group of PFs. LTD little affects PF-PC EPSPs mediated by other PFs (right). Adapted from (Tabata and Kano, 2007). (b) LTD of PF-PC EPSP induced by conjunctive PF and CF stimulation (1 Hz for 5 min). Trace: sample field EPSPs recorded before and 30 min after the conjunctive stimuli (‘‘CJS’’) are superimposed. Plot: the amplitude of field EPSP as a function of time. Adapted from (Ichise et al., 2000)

Each CF forms numerous synapses on a PC. Thus, inputs from a CF evoke a strong depolarization accompanied by action potentials and open VGCCs in the PC (> Figure 6-6). Ca2+ entering through the VGCCs activates PKC in cooperation with Ca2+ released from the ER and DAG. A possible target of PKG and PKC is a signaling cascade involving mitogen-activated protein kinase (MEK1/2) and extracellular signal-regulated kinase (ERK1/2) (> Figure 6-6). Another possible target of PKC is a nonreceptor protein tyrosine kinase(s). These possible targets are shown to be required for cerebellar LTD (Boxall et al., 1996; Kawasaki et al., 1999). There are many other signaling molecules required for cerebellar LTD although their roles in LTD induction are unclear. Such molecules include corticotrophin-releasing factor (CRF) and its receptor

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. Figure 6-6 Signaling cascades involved in cerebellar LTD induction. Signaling molecules that are shown to be required for cerebellar LTD and their possible interactions are depicted schematically. The process of the down-regulation of postsynaptic AMPAR is shown in > Figure 6-8 and omitted from this diagram. See the text for the definition of the abbreviations and further explanation. Adapted from (Tabata and Kano, 2007)

(CRF1R), Clbn1, 2-AG and CB1R, and a-calcium/calmodulin-dependent protein kinase II (a-CaMKII) (Miyata et al., 1999; Hirai et al., 2005b; Safo and Regehr, 2005; Hansel et al., 2006). CRF is produced by inferior olive neurons, and its possible target is CRF1R in PCs. Cbln1 is a cerebellum-specific protein belonging to a family including C1q complementary factor and tumor necrosis factor. Cbln1 is produced and released by GCs. 2-AG/CB1R-mediated signaling (> Section 3.3) could facilitate NO production. a-CaMKII is expressed only in PCs in the cerebellum and thus, should act postsynaptically. The final step of cerebellar LTD is the down-regulation of postsynaptic AMPAR (Matsuda et al., 2000; Linden, 2001) (> Figure 6-8). An AMPAR is a tetrameric assembly of subunits termed GluR1-4 in various combinations. PCs express GluR2, GluR3, and GluR4c. These subunits contain PSD-95/discs large/zona occludens-1 (PDZ) ligands at their C-termini. Most AMPARs in PCs include GluR2. Unless phosphorylated, the PDZ ligand of GluR2

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. Figure 6-7 Gi/o protein-coupled receptors modulate mGluR1 signaling which plays a central role in cerebellar LTD induction. GABABR enhances mGluR1 signaling through a Gi/o protein-independent pathway in response to cerebrospinal levels of GABA and Ca2+ and through a Gi/o protein-dependent pathway in response to a higher level of GABA spilt over from neighboring inhibitory synapses. These enhancements might increase the inducibility of cerebellar LTD. A1R exerts a Gi/o-independent inhibitory effect on mGluR1 signaling in response to a cerebrospinal level of adenosine. When the dose of adenosine is experimentally raised to a higher level, A1R exerts a Gi/o-dependent enhancing effect on mGluR1 signaling

binds to the PDZ domain of glutamate receptor-interacting protein (GRIP) (Matsuda et al., 2000) (> Figure 6-8a). GRIP anchors GluR2-containing AMPAR to the postsynaptic density (PSD) and thereby stabilizes its postsynaptic expression. Protein phosphatase 2A (PP-2A) dephosphorylates the PDZ ligand and can promote the postsynaptic expression of GluR2-containing AMPAR (Launey et al., 2004) (> Figure 6-8b). By contrast, PKC and MEK phosphorylate the PDZ ligand and can promote the endocytosis of GluR2-containing AMPAR (Tatsukawa et al., 2006). These phosphatase and kinases have weak basal activities; a balance between these basal activities may determine the density of AMPARs at the postsynaptic site in the absence of LTD-inducing stimuli (Launey et al., 2004; Tatsukawa et al., 2006). In response to LTD-inducing stimuli, PKC strongly phosphorylates the PDZ ligand of GluR2 (> Figure 6-8b). PKG could enhance this phosphorylation by inhibiting PP-2A via G-substrate (Launey et al., 2004). Phosphorylated GluR2 is detached from GRIP and then binds to protein interacting with C-kinase 1 (PICK1), another PDZ domain-containing protein (Xia et al., 2000). An alternative possibility is that GluR2 binds to PICK1 and then PICK1 facilitates the phosphorylation of GluR2. PICK1 forms complexes with GluRd2, a member of the ionotropic glutamate receptor family (Yawata et al., 2006). GluRd2 is shown to be required for cerebellar LTD (Kashiwabuchi et al., 1995; Hirai et al., 2003). GluRd2 does not appear to contribute to cerebellar LTD through its ligand-sensitivity or ion-permeability (Hirai et al., 2005a; Kakegawa et al., 2007). GluRd2, together with PICK1, might promote the internalization of AMPAR (Yawata et al., 2006). It is also reported that GluRd2 itself is phosphorylated during LTD induction (Kondo et al., 2005). AMPAR is finally internalized by clathrin-dependent endocytosis (Wang and Linden, 2000) (> Figure 6-8c, d). The NO-PKG cascade may facilitate endocytosis via the MEK1/2-ERK1/2 cascade (Endo and Launey, 2003a; Endo and Launey, 2003b). Genetic knock-out of GluR2 abolishes cerebellar LTD despite the continued expression of other PDZ ligand-containing GluRs (i.e., GluR3 and GluR4c) (Chung et al., 2003). This indicates that in addition to

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. Figure 6-8 Possible mechanisms underlying the final step of cerebellar LTD induction. (a) In the dendritic spine of a PC, GluR2-containing AMPAR is anchored to PSD via GRIP, and this stabilizes the postsynaptic expression of the receptor. (b) Intracellular signaling triggered by LTD-inducing stimuli promotes the phosphorylation of GluR2 (‘‘P’’), the unbinding of GluR2 from GRIP, and the binding of GluR2 to PICK1. (c) These molecular reactions facilitate the clathrin-dependent endocytosis of AMPAR. (d, e) Internalized AMPAR could be recycled. (e, a) NSFGluR2 interaction promotes the entry of GluR2-containing AMPAR into the postsynaptic site and the receptor’s competences to undergo LTD. GluR2-free AMPARs are omitted from this scheme for simplicity. Adapted from (Tabata and Kano, 2007)

the PDZ ligand, another motif peculiar to GluR2 is necessary for cerebellar LTD. A candidate for such a motif is the NSF-binding site of GluR2. An in vitro study shows that NSF-GluR2 interaction facilitates unbinding of PICK1 from GluR2 (Hanley et al., 2002). NSF-GluR2 interaction may be required for the entry of surface-expressed GluR2-containing AMPAR (> Figure 6-8e) into the postsynaptic site (> Figure 6-8a) and for the receptor’s competence to undergo LTD (Steinberg et al., 2004).

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Plasticity of CF-PC Synapses

4.1 Paired-Pulse Depression (PPD) at CF-PC Synapses When a pair of electrical pulses is given to a CF with a short interval (50–3,000 ms), CF-PC EPSC evoked by the 2nd pulse is reduced as compared with that evoked by the 1st pulse (Hashimoto and Kano, 1998) (> Figure 6-2b). Such short-lived depression is called PPD. PPD at CF-PC synapses is largely due to a usedependent decrease of glutamate release by the CF. PPD can be used as a measure to monitor the modulation of the presynaptic release machinery.

4.2 Depolarization-induced suppression of excitation (DSE) at CF-PC Synapses A prolonged depolarizing stimulus to a PC results in the production and release of endocannabinoids, which mediate DSE at CF-PC synapses (see > Section 3.3).

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4.3 LTD at CF-PC Synapses Low-frequency stimuli (e.g., 5 Hz for 30 s) to a CF innervating a PC whose membrane potential is monitored under current clamp (i.e., the PC can freely discharge) induce LTD of CF-PC EPSC lasting over 40 min (Hansel and Linden, 2000). This form of LTD requires an increase in the [Ca2+]i and the activation of mGluR1 and PKC in the PC. This form of LTD is due to a change in the glutamate-sensitivity of the postsynaptic membrane. This form of LTD may influence the inducibility of cerebellar LTD (> Section 3.5).

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Plasticity of IN-PC Synapses

5.1 STP at interneuron (IN)-PC Synapses Low-frequency CF stimulation (e.g., 2 Hz for 5 s) induces heterosynaptically STP (13 min) of inhibitory postsynaptic current (IPSC) at IN-PCsynapses (Duguid and Smart, 2004). CF-PC EPSPs open VGCCs in the PC. Ca2+ entry through the VGCCs might trigger glutamate release by the PCs (Duguid and Smart, 2004). Glutamate might bind retrogradely to NMDAR in the INs. Ca2+ entering through NMDAR might trigger Ca2+-induced Ca2+ release from the ER (see Iino, 1990 for review), and this might enhance GABA release by the INs (Duguid and Smart, 2004).

5.2 STD at IN-PC Synapses High-frequency CF stimulation (e.g., 10–40 pulses at 50 Hz) induces heterosynaptically STD (5–10 s) of IPSC at BC-PC IPCS synapses (Satake et al., 2000). This form of STD may be due to the AMPAR-mediated modulation of the presynaptic release machinery (Satake et al., 2000). Glutamate may spill over from the CF-PC synapses during the high-frequency stimuli and activate AMPAR at the presynaptic terminals of BCs (Satake et al., 2006).

5.3 LTP at IN-PC Synapses: Rebound Potentiation (RP) Low-frequency CF stimulation (e.g., 5 pulses at 0.5 Hz) induces heterosynaptically LTP (over 1 h) of IN-PC IPSC (Kano et al., 1992) (> Figure 6-9a). This form of LTP is termed rebound potentiation (RP). RP is due to an increase in the GABA-sensitivity of the postsynaptic membrane (Kano et al., 1992). RP requires an increase in the [Ca2+]i in the PC (Kano et al., 1992) (> Figure 6-9b). This may trigger the activation of CaMKII, which in turn enhances the activity of GABAAR (Kawaguchi and Hirano, 2002). RP also depends on PKA. Co-stimulation of INs disrupts RP induction presumably by activating GABABR which inhibits PKA via Gi/o protein (Kawaguchi and Hirano, 2000) (> Figure 6-9b). RP may prevent the over-excitation of PCs by CF inputs and influence motor learning.

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Plasticity of PF-IN Synapses

6.1 STD at PF-SC Synapses High-frequency PF stimulation (e.g., 10 pulses at 50 Hz) induces STD of PF-SC EPSC lasting for 20 s (Beierlein and Regehr, 2006). This form of STD requires 2-AG/CB1R-mediated retrograde signaling (> Section 3.3) (Beierlein and Regehr, 2006). In SCs, the activation of mGluR1 and NMDAR may trigger the production of 2-AG (Beierlein and Regehr, 2006).

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. Figure 6-9 RP (LTP at IN–PC synapses). (a) RP induced by low-frequency CF stimulation (5 pulses at 5 Hz). In this example, spontaneous IPSCs were measured (traces) and their mean amplitude for each 2-min period is plotted as a function of time after the CF stimuli. Adapted from (Kano et al., 1992). (b) Possible mechanisms underlying RP (left side of this panel). See the text for further explanation. Co-stimulation of INs results in GABABR activation in the PC, and this prevents RP induction (right side of this panel). CaM, calmodulin; PP-1, protein phosphatase-1; AC, adenylyl cyclase; D-32, DARPP-32. Adapted from (Kawaguchi and Hirano, 2002)

This form of STD may occur in vivo when PFs discharge at a high rate (see > Section 3.2). This form of STD may transiently relieve GC-SC-PC feedforward inhibition (> Section 2). The relief of the feedforward inhibition extends the time-window for PF-carried signals to contribute to information integration in PCs; increases the efficacy of PC-DCNN synaptic transmission; and may influence various forms of synaptic plasticity involving PCs.

6.2 LTP/LTD at PF-SC Synapses Low-frequency PF stimulation (e.g., 2 Hz for 60 s) induces LTP or LTD of PF-SC EPSC lasting over 30 min (Rancillac and Crepel, 2004). The probability of occurrence is almost equal for LTP and LTD. LTP of PF-SC EPSC requires NO and the activation of group III mGluRs (Rancillac and Crepel, 2004) LTD of PF-SC EPSC requires the activation of group II mGluRs in the SC (Rancillac and Crepel, 2004). When combined with a depolarizing stimulus (e.g., 0 mV) to the SC, the low-frequency PF stimulation mostly induces LTP of PF-SC EPSC lasting over 30 min (Rancillac and Crepel, 2004). This form of LTP requires an increase in the [Ca2+]i in the SC, the activation of NMDAR, and NO (Rancillac and Crepel, 2004). Temporally correlated activities of PFs and a CF that induce cerebellar LTD may also induce this form of LTP, because CFs project their Scheibel collaterals to SCs. Thus, this form of LTP could play a role in cerebellar LTD-dependent motor learning. Moderately high-frequency PF stimulation (e.g., 8 Hz for 15 s) induces LTP of PF-SC EPSC lasting over 30 min in a cAMP-dependent manner (Rancillac and Crepel, 2004). High-frequency burst PF stimulation (e.g., 4 sets of bursts at 0.33 Hz, each burst consists of 25 pulses at 30 Hz) induces LTD of PF-SC EPSC lasting over 30 min (Soler-Llavina and Sabatini, 2006). This form of LTD requires Ca2+ entry through AMPAR and the activation of mGluR1 and CB1R, but not Ca2+ release from the ER (Soler-Llavina and Sabatini, 2006). In addition, high-frequency PF stimulation (e.g., 300 pulses at 50 Hz) reduces the Ca2+-permeability of AMPARs in the SC over 60 min (Liu and Cull-Candy, 2000). This long-term change in receptor function may be due to the down-regulation of Ca2+-permeable AMPAR subtypes and the up-regulation of Ca2+-impermeable subtypes (Liu and Cull-Candy, 2000).

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LTP at mossy fiber (MF)-deep cerebellar nuclear neuron (DCNN) Synapses

Conjunctive pre- and postsynaptic stimuli (> Figure 6-10b, schematics) that mimic synaptic inputs received by a DCNN during delayed eye-blink conditioning (see > Section 9) (> Figure 6-10a) induce LTP of MF-DCNN EPSC lasting over 20 min (> Figure 6-10c) (Pugh and Raman, 2006). PCs discharge at a high rate at the early phase of the conditioning (see Pugh and Raman, 2006 for review), and their inhibitory synaptic outputs may hyperpolarize the DCNNs transiently. Upon the termination of a hyperpolarization, DCNNs display a rebound depolarization (RD) (> Figure 6-10b, sample record) presumably due to inward conductance through low-threshold VGCC, hyperpolarization-activated cation channel, and Na+ channels (Pugh and Raman, 2006). A RD opens VGCCs. On the other hand, glutamate released by the MFs activates NMDAR in the DCNN. Ca2+ entry through both the VGCCs and NMDAR is required for this form of LTP (Pugh and Raman, 2006).

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LTD at PC-DCNN Synapses

High-frequency burst CF stimulation (e.g., 10 sets of burst at 2 Hz, each burst consists of 10 pulses at 100 Hz) induces LTD of PC-DCNN IPSP lasting over 20 min (Aizenman et al., 1998). Each burst evokes a sustained hyperpolarization, which is followed by RD (> Section 7) in the DCNN (Aizenman et al., 1998). When RD elicits more than five action potentials, a high level of Ca2+ enters through VGCCs, and this triggers LTD induction (Aizenman et al., 1998).

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Physiological Significance of Cerebellar Synaptic Plasticity

9.1 Modern Behavioral Studies Classical behavioral studies employing cerebellar lesions indicate the importance of the cerebellum for motor coordination, motor accuracy, and motor learning. Researchers are now trying to elucidate the behavioral-level role of specific forms of cerebellar synaptic plasticity by pharmacologically and/or genetically modulating the underlying mechanisms. In the following sections, we review recent findings obtained through such an approach.

9.2 Motor Coordination When a normal mouse walks on a treadmill spinning at a fixed speed, the step cycles of the left and right fore-/hindlimbs alternate regularly with phase differences close to 180 degrees (Ichise et al., 2000) (> Figure 6-11a). The mGluR1-KO mice (> Section 3.5.2) display irregular alternation with more dispersed phase differences, indicating the loss of interlimb coordination (Ichise et al., 2000). PC-specific genetic rescue of mGluR1 in the mGluR1-KO mice (mGluR1-rescue mice, > Section 3.5.2) restores regular alternation (Ichise et al., 2000). An untrained mouse can remain on a rotating bar for only a few s (rota-rod task). However, a normal mouse is able to learn how to move its limbs to remain on the rotating bar through training sessions, and retention time is extended trial after trial (> Figure 6-11b). Learning in this task is severely impaired in the mGluR1-KO mice, whereas partially restored in the mGluR1-rescue mice (Ichise et al., 2000). These observations suggest the importance of cerebellar LTD for the execution and learning of motor coordination. However, glial fibrillary acidic protein (GFAP)-KO mice (Shibuki et al., 1996) and the CB1R-KO mice (Kishimoto and Kano, 2006) display normal learning in rota-rod task despite their deficiency in cerebellar LTD. The precise role of cerebellar LTD in motor coordination is to be elucidated in future.

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. Figure 6-10 LTP at MF-DCNN synapses. (a) Diagram of the predicted firing patterns of the labeled fibers and neurons during delayed eye-blink conditioning (cf. > Figure 6-12). ‘‘+’’ and ‘‘’’: excitatory and inhibitory synapses, respectively. (b) Conjunctive pre- and postsynaptic stimuli for mimicking synaptic inputs during delayed eye-blink conditioning (schematics) and a sample voltage response of a DCNN to these stimuli (trace). The conjunctive stimuli consist of a high-frequency pulse train (133 Hz) delivered to the MFs and a hyperpolarizing current (300–500 pA) delivered to the DCNN under current clamp. (C) LTP of MF-DCNN EPSC induced by the above conjunctive stimuli. The recordings were made under voltage clamp except the period of the conjunctive stimulation. Plot: the amplitude of EPSC recorded from a DCNN as a function of time after the conjunctive stimuli. Open and closed symbols: individual data and the averages of 10 measurements, respectively. Traces: sample EPSCs recorded before and after the conjunctive stimuli. All panels adapted from (Pugh and Raman, 2006)

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. Figure 6-11 Motor coordination depends on cerebellar LTD. (a) Genetic knock-out of mGluR1 [‘‘mGluR1(2/2)’’] impairs interlimb coordination during walk whereas PC-specific genetic rescue of mGluR1 (‘‘mGluR1-rescue’’) restores it. Plots: the histograms of phase differences between the step cycles of left and right fore-/hindlimbs of the labeled mice walking on a treadmill spinning at a fixed speed. (b) Genetic knock-out of mGluR1 abolishes learning in rota-rod task (i.e., trial-after-trial increase in performance) whereas PC-specific genetic rescue of mGluR1 restores it. Performance was evaluated as the time for which the mouse remained on a rotating bar. The extent of restoration depends on the expression level of mGluR1 (‘‘mGluR1-rescue Tg/Tg’’ mice that are homozygous for the transgene express a higher level of mGluR1 than ‘‘mGluR1-rescue’’ mice that are heterozygous for the transgene). All panels adapted from (Ichise et al., 2000)

9.3 Delayed Eye-blink Conditioning Eye-blink reflex is an inherited reflex evoked by aversive periorbital stimuli (e.g., air puff to the eye or an electrical shock to the eyelid; unconditioned stimuli, US) (> Figure 6-12a). This reflex becomes responsive to behaviorally neutral stimuli (conditioned stimuli, CS; e.g., pure tone) after the animal is exposed to CS paired with US for several times (eye-blink conditioning). In the delayed version of this conditioning, a CS starts prior to and co-terminates with an US. Learning in delayed eye-blink conditioning is dependent on the cerebellum (Linden, 2003; De Zeeuw and Yeo, 2005). The information of US and CS is conveyed to PCs by CFs and PFs, respectively. Learning in delayed eye-blink conditioning is impaired in the mGluR1-KO mice (> Section 3.5.2) (Aiba et al., 1994, Kishimoto et al., 2002) (> Figure 6-12b), the GFAP-KO mice (Shibuki et al., 1996), GluRd2-KO mice (Kishimoto et al., 2001), transgenic mice with PCs expressing a PKC inhibitor peptide under the control of L7 promoter (L7-PKCi mice) (Koekkoek et al., 2003), and the CB1R-KO mice (Kishimoto and Kano, 2006). These mouse strains are commonly deficient in cerebellar LTD. Moreover, normal learning is

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. Figure 6-12 Delayed eye-blink conditioning depends on cerebellar LTD. (a) Neural circuits involved in delayed eye-blink conditioning. ‘‘+’’ and ‘‘2’’: excitatory and inhibitory synaptic transmission, respectively. Adapted from (Linden, 2003). (b) Genetic knock-out of mGluR1 [‘‘mGluR1(2/2)’’] attenuates a CS-evoked eye-blink response (CR) whereas PC-specific genetic rescue of mGluR1 restores it. Schematics: the timing of a CS (tone) and a US (electrical shock to the eyelid) presented for training. Plots: CRs of the labeled mice tested on the seventh day of daily training sessions. CRs were measured as electromyograms (EMG) recorded from the eyelids. Adapted from (Kishimoto et al., 2002)

seen in the mGluR1-rescue mice in which cerebellar LTD can be induced (> Section 3.5.2) (Kishimoto et al., 2002). These observations together suggest the involvement of cerebellar LTD. Pharmacological and morphological studies suggest that synaptic plasticity in the DCN is also involved in learning in delayed eye-blink conditioning (see DeZeeuw and Yeo, 2005 for review). Conjunctive MF and DCNN stimulation that mimics synaptic inputs received by a DCNN during delayed eye-blink conditioning induce LTP of MF-DCNN EPSC (see > Section 7).

Synaptic plasticity in the cerebellum

6

9.4 Adaptation of Vestibulo-Ocular Reflex (VOR) and Optokinetic Reflex (OKR) VOR is compensatory eye movement to minimize the blur of the visual scene due to head movement. When an untrained animal in motion (e.g., an animal mounted on a turntable rotating left and right) is exposed to the visual scene (e.g., a patterned screen swinging left and right, > Figure 6-13a) moving to a smaller or larger extent than expected from head movement (visual-vestibular mismatch task), the eyes cannot precisely track the visual scene. However, tracking fidelity (VOR gain) is improved gradually through training sessions (VOR adaptation). The cerebellar flocculus, an evolutionally old part of the cerebellar cortex, plays an important role in VOR adaptation (> Figure 6-13a). The sensory information of head/eye movement and the visual informationof visual scene blur are conveyed to PCs in the flocculus by PFs and CFs, respectively. Cerebellar LTD triggered by CF inputs modifies the manner of information integration in and the outputs of the PCs. This change in the outputs may influence eye movement regulation by vestibular nuclear neurons. Pharmacological inhibition of cerebellar glutamate receptors during or the flocculus immediately after a training session cancels VOR adaptation (see Gittis and DuLac, 2006 for review). The L7-PKCi mice do not show VOR adaptation after short-term training (1 h) (DeZeeuw and Yeo, 2005). These observations together suggest that cerebellar LTD in the flocculus is important for at least the early stage and/or short-term memory of VOR adaptation. Some in vivo studies indicate that VOR learning is accompanied by synaptic plasticity in the vestibular nuclei (see Gittis and DuLac, 2006 for review). LTD occurs at synapses from PCs to some postsynaptic neurons (> Section 8). Thus, the vestibular nuclei may be involved in VOR adaptation. OKR is compensatory eye movement to minimize the blur of the visual scene due to the movement of the visual scene itself. The tracking fidelity of the eyes to the visual scene (OKR gain) is improved gradually through training sessions (> Figure 6-13b). The neural circuit involved in OKR largely overlaps that in VOR (> Figure 6-13a). Optokinetic signals are transmitted from the accessory optic system to the cerebellar flocculus and the vestibular nuclei. Pharmacological inhibition of the flocculus erases the short-term memory of OKR adaptation acquired in a training session on the same day, but not the long-term memory acquired in training sessions on the previous days (Shutoh et al., 2006) (> Figure 6-13c). Pharmacological inhibition or genetic knock-out of neuronal NO synthase that is required for cerebellar LTD impairs OKR adaptation (Shutoh et al., 2006). These observations together suggest that cerebellar LTD in the flocculus is important for the early stage or short-term memory of OKR adaptation, while another brain region(s) retains the long-term memory. Vestibular nerve-vestibular nuclear neuron synapses are potentiated after daily training sessions for 1 week (Shutoh et al., 2006), suggesting that synaptic plasticity in the vestibular nuclei is involved in the retention of the long-term memory.

9.5 Fear Conditioning Animals such as rats freeze in response to fearful stimuli (US; e.g., electrical shocks from the floor). After trained with a few pairs of pure tones (CS) and fearful stimuli (auditory fear conditioning), an animal freezes in response to test presentations of pure tones alone. Once trained, an animal retains the memory of auditory fear conditioning for several days. An animal also learns the context in which fearful stimuli are presented (i.e., environmental clues in the conditioning apparatus) (contextual fear conditioning) and become responsive to a test presentation of the context alone. To assess the auditory component of learning in fear conditioning, test pure tones are presented in an apparatus different than that used for conditioning. To assess the contextual component, a trained animal is placed in the conditioning apparatus. For both components, learning is typically evaluated as the total time of freezing during a test presentation(s). It is widely accepted that amygdala is important for fear memory. A study (Sacchetti et al., 2002) revealed that the cerebellum is also important for fear memory. Reversible pharmacological inactivation of

81

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Synaptic plasticity in the cerebellum

. Figure 6-13 VOR and OKR adaptation depends on cerebellar synaptic plasticity. (a) Neural circuit involved in VOR and OKR. Arrows: the flows of information of head movement necessary for VOR and optikinetic signals necessary for OKR. ‘‘STM’’ and ‘‘LTM’’: the possible locations of the short-term and long-term memory traces of OKR adaptation, respectively. AOT: accessory optic tract; CF: climbing fiber; FL: cerebellar flocculus; GC: granule cell; IO: inferior olive; OMN: oculomotor neuron; PC: Purkinje cell; PF: parallel fiber; VN: vestibular nucleus; VO: vestibular organ. (b) OKR gain increases gradually through daily training sessions. Open and closed dots: OKR gains measured before and after a training session on the day, respectively. Gray dots: OKR gain after the termination of the training sessions. Inset: the movements of the patterned screen and the eye on the labeled day of daily training sessions. (c) Injection of lidocaine, an anesthetic into the cerebellar flocculus on the fourth day of daily training sessions erases the short-term memory of OKR adaptation acquired in a training session on the same day [compare OKR gains measured after a training session on the fourth day (right middle bar) and 30 min after lidocaine injection (right most bar)]. However, this manipulation little affects the longterm memory acquired in training sessions from the first to third days [note that lidocaine does not lower OKR gain below the level measured before a training session on the fourth day (left middle bar)]. All panels adapted from (Shutoh et al., 2006)

the vermis (the midline part of the cerebellar cortex) after a training session impairs both the auditory and contextual components of learning in fear conditioning. Reversible pharmacological inactivation of the interposed nuclei after a training session impairs the auditory, but not the contextual component. Drug administration even with a very long delay (up to 96–192 h) after a training session is effective. These observations suggest that the vermis and interposed nuclei are involved in the consolidation of fear memory.

Synaptic plasticity in the cerebellum

6

In the lobules V–VI of the vermis in slice preparations prepared 10 min or 24 h after a training session of auditory fear conditioning, PF-PC EPSCs are potentiated (Sacchetti et al., 2004). This potentiation may be due to an increased postsynaptic expression of AMPAR , but not an increase of glutamate release from the presynaptic membrane (Sacchetti et al., 2004). Such potentiation is not seen for CF-PC EPSCs (Sacchetti et al., 2004). These observations suggest that a form of LTP of PF-PC EPSC is involved in the consolidation of fear memory.

10

Conclusive Remarks

Since the dawn of cerebellar synaptic plasticity research, LTD at PF-PC synapses has been highlighted as a principal physiological basis for motor learning, and investigated extensively at both the molecular and behavioral levels. However, it was later shown that virtually all of the cerebellar synapses undergo various forms of synaptic plasticity. For most of these forms of synaptic plasticity, the underlying molecular mechanisms and physiological significance remain to be explored. As demonstrated by the studies using the mGluR1-rescue and L7-PKCi mice, cell type-specific genetic manipulation would be a powerful method to analyze each specific form of cerebellar synaptic plasticity. Moreover, recent studies reveal the involvement of cerebellar synaptic plasticity in fear memory. Therefore, the range of exploration should be expanded to non-motor functions.

Acknowledgments Our work cited in this chapter was partly supported by Grants-in-Aid for Scientific Research from the Ministry of Education, Science, Sports, Culture and Technology of Japan (17,700,305; 18,019,022; 19,045,019; 20,022,025; and 20,500,284 to T.T., 17,023,021 and 17,100,004 to M.K.).

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7

Vesicular Neurotransmitter Transporters

H. Fei . D. E. Krantz

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89

2

History of Identification and Cloning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89

3

Phylogeny . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91

4 4.1 4.2 4.3 4.4 4.5 4.6

Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 92 Bioenergetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 92 Neurotransmitter Content . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Variation in Quantal Size . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94 Rate of Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 When are Vesicles Loaded? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Transporter Substrates and Their Affinities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96

5

Pharmacology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97

6 6.1 6.2 6.3

Models and Structure/Function Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 Membrane Topology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 Structure/Function Studies of Transport Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99

7 7.1 7.2 7.3 7.4 7.5 7.6

Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102 The CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102 Glia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Mismatched Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 Outside the CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 Developmental Changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 Co-Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105

8 8.1 8.2 8.3

Biosynthesis and Trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Membrane Trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 How Many Transporters are on Each Vesicle? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 110

9 9.1 9.2 9.3 9.4 9.5 9.6

Regulation of Expression and Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Transcription . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 Alternative mRNA Splicing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Trafficking and Transporter Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Heterotrimeric G Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 114 Protein Binding Partners (see also Section 8.2) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Upstream Regulatory Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115

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10 Functional Effects of Altered Expression and Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 10.1 Regulation of Quantal Size . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 116 10.2 Cytoplasmic Clearance of Dopamine and Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 11 11.1 11.2 11.3 11.4

Behavioral Genetics of Animal Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 Mouse Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 C. elegans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 Drosophila . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 Zebrafish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121

12 Human Genetic Studies and Disease Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121 12.1 Genetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121 12.2 Altered Expression in Human Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122 13

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122

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Abstract: Vesicular transporters fill synaptic and other types of secretory vesicles with neurotransmitter. This structurally divergent family includes specific transporters for the monoamines (VMATs), acetylcholine (VAChT), glutamate (VGLUTs) and GABA+glycine (VGAT or VIAAT). We review the pharmacology, expression, membrane trafficking, and regulation of each subtype. We also discuss the behavioral effects of genetic mutations and knockouts, and the role that vesicular transporters may play in controlling the amount of neurotransmitter that is released from each vesicle. List of Abbreviations: ATP, adenosine triphosphate; DA, dopamine; ECL, enterochromaffin-like; Epi, epinephrine/adrenalin; 5HT, serotonin; GABA, gamma amino butyric acid; LDCG, large dense core granule; LDCV, large dense core vesicle; MDMA, 3,4 methylenedioxymethamphetamine; MFS, major facilator superfamily; MPP+, N-methyl-4-phenylpyridium; NE, norepinephrine/noradrenalin; SIF, small intensely fluorescent; SV, synaptic vesicle; TM, transmembrane domain; VAChT, vesicular acetylcholine transporter; VGAT, vesicular GABA transporter; VGLUT, vesicular glutamate transporter; VIAAT, vesicular inhibitory amino acid transporter

1

Introduction

Secretory vesicles release all classical and amino acid neurotransmitters. These include acetylcholine, monoamines, GABA, glycine and glutamate. However, with the exception of noradrenalin, enzymes required for their synthesis reside in the cytoplasm. Therefore, their exocytotic release requires transport out of the cytoplasm and into the vesicular lumen. This process is mediated by a class of proteins known as vesicular neurotransmitter transporters. Biochemical assays and the more recent analysis of molecularly cloned transporters have identified four families that serve to transport and store most known neurotransmitters. These include vesicular transporters for: (1) acetylcholine (the vesicular acetylcholine transporter or VAChT); (2) monoamines such as dopamine and serotonin (vesicular monoamine transporters or VMATs); (3) GABA and glycine (the vesicular GABA transporter, VGAT, also known as the vesicular inhibitory amino acid transporter or VIAAT); and (4) glutamate (vesicular glutamate transporters or VGLUTs). Additional small molecules that function as neurotransmitters such as ATP, proline and aspartate may use other vesicular transporters, but these remain to be identified. Larger peptide hormones/neurotransmitters such as endogenous opiates also undergo exocytotic release, but do not employ vesicular transporters to enter secretory vesicles. Rather, they are loaded into the lumen of large dense core vesicles (LDCVs) as secreted, precursor proteins (Kelly, 1993; Dikeakos and Reudelhuber, 2007). Some gases such as nitrous oxide (NO) also function as neurotransmitters, but these are thought to freely diffuse across biological membranes and do not require either vesicular transport or exocytotic release (Boehning and Snyder, 2003). We therefore confine our discussion to neurotransmitters acetylcholine, monoamines, GABA, glycine, and glutamate and discuss the vesicular transporters VAChT, the VMATs, VGAT and the VGLUTs (> Table 7-1).

2

History of Identification and Cloning

Active transport of neurotransmitter into vesicles was reported as early as 1962 by Arvid Carlson, among others (Carlsson et al., 1962; Kirschner, 1962). At that time, neither GABA nor glutamate had been clearly defined as neurotransmitters, and the biochemical purification of synaptic vesicles (SVs) was in its infancy. Therefore, the first vesicular uptake assays were performed using monoamine transmitters, and an aminergic tissue rich in secretory vesicles other than SVs. A series of biochemical experiments showed that uptake of monoamines into adrenal chromaffin granules is ATP dependent, and over the next twenty years the bioenergetics of transport were established using similar assays (Schuldiner et al., 1978, 1995; Johnson, 1988). The electrochemical gradients that drive vesicular transport are described in more detail below. Expression cloning strategies identified the first cDNAs encoding a VMAT (Erickson et al., 1992; Liu et al., 1992b). One group, led by Robert Edwards, screened a cDNA library from adrenal pheochromocytoma cells

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. Table 7-1 Summary of vesicular transporter subtypes Transporter VMAT1

VMAT2

VAChT VGAT/ VIAAT VGLUT1 VGLUT2 VGLUT3

Substrates Serotonin, Dopamine, Epinephrine Norepinephrine (Histamine)a Serotonin, Dopamine, Epineprhine Norepinephrine, Histamine Acetylcholine GABA, glycine Glutamate Glutamate Glutamate

Apparent affinity (cell free assay) 0.85 1.56 1.86 2.5 (436) 0.19 mM 0.32 0.47 0.33 3.06 1 mM 5 mM 25 mM 3.4 mM 0.8 mM 0.5 mM

Phylogenetic classification SLC18A1

prior names CGAT

C.elegans ortholog

Drosophila ortholog

SLC18A2

SVAT

cat-1

dVMAT

unc-17 unc-47

dVAChT dVGAT

eat-4

dVGLUT

SLC18A3 SLC32 SLC17A6 SLC17A7 SLC17A8

BNPI DNPI

Some of the properties of vesicular neurotransmitter transporters are summarized here. See text for additional details a Histamine is not usually considered a high affinity substrate for VMAT1

(Liu et al., 1992a, 1992b). By expressing the library in an epithelial cell line that was sensitive to the neurotoxin N-methyl-4-phenylpyridium (MPP+), they identified a cDNA that conferred resistance to the toxin; they later showed it encoded a transporter that sequestered MPP+ into vesicles and away from its site of action in mitochondria. Originally named the chromaffin granule amine transporter (CGAT) it was later renamed VMAT1. A homology-based cloning strategy was used to identify a second rat VMAT isoform expressed in the brain that was eventually designated VMAT2. Parallel experiments by Jeff Erickson, Lee Eiden and Beth Hoffman independently identified VMAT2 directly - also using an expression cloning strategy (Erickson et al., 1992). A third group led by Shimon Schuldiner used affinity purification to identify VMAT2 (SternBach et al., 1992). In each case, the cDNA clones representing both VMAT1 and 2 were found to confer robust transport activity in vitro, and their expression in aminergic cells confirmed that they indeed represented vesicular monoamine transporters. Similar to the importance of the adrenal gland in the study of monoamines, the electronic organ tissue of the Torpedo fish provided a critical tissue source for early studies on acetylcholine storage. It too, was found to be dependent on an electrochemical gradient and over the past 25 years, a series of detailed and elegant experiments on VAChT by Stan Parsons’ lab and others have helped determine the mechanisms underlying acetylcholine transport (Parsons, 2000). The drug vesamicol played an important role in these experiments and VAChT is indicated as the vesamicol binding protein in some papers published prior to its molecular identification. Although much of the biochemistry of acetylcholine transport was established using tissue from Torpedo, studies in C. elegans were critical to identifying the gene encoding VAChT. Genetic screens for uncoordinated mutants first identified the unc-17 gene, later shown to map within the choline acetyltransferase (chat) gene responsible for acetylcholine synthesis (Rand and Russell, 1984). Biochemical studies showed that acetylcholine levels were altered in unc-17 mutants, thus suggesting a defect in acetylcholine storage and/or transport (Hosono et al., 1987). Since unc-17 was found to be less sensitive to blockade of acetylcholine breakdown, and appeared similar in structure to VMAT, it was suggested that it might encode a Caenorhabdlitis elegans (C. elegans) ortholog of VAChT (Alfonso et al., 1993). Based on the similarity of unc-17 to VMATs, oligonucleotides representing unc-17 were used to identify potential

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orthologs of VAChT from Torpedo, rat and human (Erickson et al., 1994; Roghani et al., 1994; Varoqui et al., 1994). Biochemical experiments in the Erickson lab finally confirmed that the rVAChT cDNA conferred acetylcholine uptake activity in vitro (Varoqui and Erickson, 1996). Meanwhile, the idea that VaChT could function as true vesicular transporters was aided by electrophysiological experiments using frog co- cultures of cholinergic neurons and myocytes (Song et al., 1997). The notion that amino acids such as GABA and glutamate could function as neurotransmitters was not fully accepted until the 1970s, and vesicular GABA uptake was not demonstrated until the late 1980s, using synaptic vesicle preparations from mammalian tissue (Fykse and Fonnum, 1988; Hell et al., 1988; Kish et al., 1989). Similar to VAChT, the cloning of VGAT/VIAAT was based on studies originally performed in C. elegans. Ablation experiments had identified GABAergic neurons in the worm, and subsequent studies showed that mutation of the predicted gene, unc-47, caused a phenotype similar to ablation of the GABAergic neurons (McIntire, 1993). Molecular cloning of unc-47 revealed that it was structurally similar to amino acid transporters and thus likely to represent a transporter (McIntire et al., 1997). Homology based cloning strategies were then used by the Edwards and Gasnier labs to isolate a mammalian VGAT/ VIAAT homolog which displayed robust GABA transport activity in vitro, similar to activities previously found in brain tissue (McIntire et al., 1997; Sagne et al., 1997). Vesicular glutamate transport was characterized using synaptic vesicle preparations similar to those used for GABA uptake (Disbrow et al., 1982; Naito and Ueda, 1983). However, the VGLUTS were the last of the four known vesicular transport activities to be molecularly cloned. Some delay was caused by the hypothesis that the VGLUTs would be structurally similar to VGAT, which led to homology-based searches for VGLUTs. Interestingly, these screens eventually led to the identification of the transporters for Systems N and A (i.e., SNATs, see > Section 3) (Chaudhry et al., 1999). Meanwhile, two novel genes in a family of plasma membrane Na+- phosphate transporters were cloned, including one gene expressed in cerebellar neurons and upregulated by the glutamate receptor agonist NMDA. It was designated the Brain Specific Na+-dependent phosphate transporter, or BNPI (Ni et al., 1994). A second gene was cloned from rat pancreatic cells and was shown to be upregulated during differentiation to a neuroendocrine phenotype (Aihara et al., 2000). The second gene was therefore named Differentiation Associated Na+-dependent phosphate transporter (DNPI). Although the original Pi uptake assays were based on the activity of BNPI and DNPI at the plasma membrane, BNPI was later shown to localize to synaptic vesicles (Bellocchio et al., 1998). To reconcile these apparently disparate properties, it was originally hypothesized that BNPI would localize to the plasma membrane during SV exocytosis and regulate phosphate-sensitive glutamate synthesis (Bellocchio et al., 1998). However, heterologous expression in cultured cells showed that it conferred vesicular glutamate uptake, and a plethora of subsequent studies have cemented its identity as a VGLUT (Reimer and Edwards, 2004). Interestingly, recent biochemical experiments have shown that VGLUTs may indeed function as both vesicular glutamate transporters and sodium dependent phosphate transporters, and that the mechanism of each may differ (Juge et al., 2006). However, the potential physiological function of phosphate transport by VGLUTs is not clear. After the first VGLUT was identified and designated VGLUT1, additional homology based cloning efforts showed that two more VGLUT genes are expressed in mammals (VGLUT 2 and 3 (Reimer and Edwards, 2004)). This diversity was not predicted based on earlier biochemical assays and has led to a number of interesting studies on the expression, subcellular localization, and regulation of the three VGLUT isoforms (see below).

3

Phylogeny

All vesicular transporters are members of the Major Facilator Superfamily (MFS). The SLC18 group of the MFS includes VMAT1 (SLC18A1), VMAT2 (SLC18A2) and VAChT (SLC18A3). The SLC18 family is distantly related to the toxin extruding antiporters (TEXANS), bacterial transporters that have been proposed as a model for the function of the VMATs (Yelin and Schuldiner, 1995). VGAT is the lone member of the SLC32 family (Gasnier, 2004). However, it belongs to a larger super-family that includes

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SLC38 and SLC36. SLC38 includes System N/System A transporters, or SNATs, responsible for recycling glutamate and glutamine (Mackenzie and Erickson, 2004). SLC36 includes proton driven transporters PAT2 and LYAAT (Sagne et al., 2001; Wreden et al., 2003; Boll et al., 2004). VGLUT is a member of SLC17, which consists of three subfamilies. One subfamily includes the three VGLUTs, SLC17A6-8, which share 74–82% identity (Reimer and Edwards, 2004). The second subfamily is comprised of four genes whose physiological functions remain unknown (SLCA1-4), although at least one member transports both inorganic and organic anions (Butterworth et al., 1995). The third subfamily of SLC17 contains only one member, sialin (SLC17A5), which is required for H+ dependent transport of sialic acid across lysosomal membranes (Mancini et al., 1989; Reimer and Edwards, 2004).

4

Transport

4.1 Bioenergetics It is well-established that the activities of VMAT and VAChT require exchange of lumenal protons for cytosolic transmitter via an antiporter mechanism, and that the intravesicular concentration of protons is generated by the vacuolar H+-ATPase (Johnson, 1988; Parsons, 2000). By extension, it is assumed that VGLUTs and VGAT employ similar mechanisms. The requirement of the vacuolar H+-ATPase for active vesicular transport has been demonstrated in part through the use of bafilomycin, a specific inhibitor of this enzyme, which blocks both the generation of the electrochemical gradient and active neurotransmitter transport (Johnson, 1988; Parsons, 2000). Additional agents important for determining the bioenergetics of vesicular transport include the proton ionophores carbonyl cyanide m-chlorophenylhydrazone (CCCP) and carbonylcyanide p-(trifluoromethoxy) phenylhydrazone (FCCP) - agents that were first characterized using mitochondria (Kessler et al., 1977). Similar to their extensively studied action at the mitochondrial membrane, CCCP and FCCP equilibrate proton concentration across the vesicular membrane, and thereby block vesicular neurotransmitter transport. The stoichiometry of the antiport mechanism has been directly studied for VMAT (Knoth et al., 1981) and VAChT (Nguyen et al., 1998) and both transporters exchange 2 protons for each molecule of neurotransmitter that is taken up into the vesicle. The proton electrochemical gradient DmH+, which drives transport across the vesicular membrane, includes both a proton concentration component DpH and an electrical component Dc. For VMATs, and VAChT, these have been determined to be  1.5 pH units and 50 mV, respectively (Johnson, 1988; Parsons, 2000). Protons and positive charge flow down the concentration and electrical gradients respectively, and both forces are coupled to the influx of neurotransmitter. It should be noted, however, that the amount of transmitter that can be loaded into vesicles is thought to depend on factors other than DmH+, such as substrate affinity, lumenal Cl, vesicle volume and intralumenal proteins (see below). Each family of vesicular transporters has been shown to differ in their relative requirement for DpH versus Dc. This has been determined by selectively reducing either DpH or Dc, primarily using pharmacologic probes such as nigericin and valinomycin. In contrast to proton ionophores, the H+/K+ exchanger nigericin can be used to reduce DpH without disrupting the electrical gradient. Conversely, valinomycin, a K+ ionophore, can be used to reduce Dc without eliminating the proton gradient. Experiments using these and other pharmacologic agents have shown that VMATs and VAChT primarily use DpH (Parsons, 2000), whereas glutamate transport uses Dc (Reimer and Edwards, 2004). VGAT has been proposed to use both DpH and Dc (McIntire et al., 1997; Gasnier, 2004). An additional dependence on chloride has been determined for VGLUT transport activity in vitro. Dependence on Cl is biphasic, such that at low concentrations, Cl facilitates, and at high concentrations it inhibits transport (Naito and Ueda, 1985). It has been presumed that Cl ions enter the lumen of SVs via a channel, but H+/Cl exchange has been proposed as an alternative mechanism (Accardi et al., 2004; Edwards, 2007). VGLUT transport activity may be directly, allosterically stimulated by Cl (Hartinger and Jahn, 1993). Transport assays performed using VGLUT alone reconstituted into proteoliposomes suggest that the effect

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of Cl is not due to interactions with any other vesicular protein (Juge et al., 2006). However, it is also possible that Cl facilitates transport by acting as a counter ion for positively charged, lumenal protons, thereby allowing a greater DpH. Arguing against this model is the finding that an increase in transport occurs even if the lumenal proton concentration is not allowed to increase (Wolosker et al., 1996). Regardless of the mechanism by which Cl facilitates VGLUT activity, it is more likely to affect the initial rate of transport rather than steady-state levels (Wolosker et al., 1996). Furthermore, effects of this phenomenon on synaptic transmission in vivo remain unclear, since alterations in presynaptic Cl do not appear to influence quantal size (Price and Trussell, 2006). The charge on transported substrate differs between vesicular transporters, but all vesicular transport activities are thought to be electrogenic, with transport causing a net flux of positive charge. The pKa of the primary amine moiety of serotonin, dopamine and noradrenalin suggest they all will have net charge of 1 at physiological pH; in addition, the observation that permanently charged species such as MPP+ are substrates of VMAT indicates that the endogenous substrates are likely to undergo transport as positively charged ions (Daniels and Reinhard, 1988; Scherman et al., 1988). The exchange of two protons for one cationic transmitter suggests an electrogenic process with a net flux of one positive charge per VMAT transport cycle. Similarly, since acetylcholine is permanently charged (+1) at physiological pH, exchange of two protons will result in a net flux of +1. It should be noted however, that the electrical properties of vesicular transport have not been measured directly. In theory, it should be possible to record electrical activity of vesicular transporters that are forced to localize to the plasma membrane; VMAT2 forced to localize to the plasma membrane of oocytes has been shown to transport tritiated substrate (Whitley et al., 2004). However, electrophysiological experiments have not been reported. Unlike vesicular monoamine and acetylcholine transport, the stoichiometries of vesicular GABA and glutamate transport remain unclear. It has been argued that the proton electrochemical gradient may be used only to establish Dc, rather than drive actual movement of protons across the vesicular membrane (Maycox et al., 1988; Carlson et al., 1989a). However, reducing Dc reveals a residual transport activity that can be abolished by reducing DpH (Tabb et al., 1992), more consistent with the notion that proton flux as well as Dc are required for vesicular glutamate transport as they are for amine and acetycholine transport.

4.2 Neurotransmitter Content The question of how much transmitter is stored in each vesicle can be estimated from the strength of the proton electrochemical gradient. Transport by the VMATs involve the exchange of two lumenal protons for one protonated molecule of transmitter, and consequent outward movement of two protons and one net positive charge. For a vesicular pH gradient of 1.5 pH units and membrane potential of 50 mV, this predicts accumulation of transmitter inside vesicles 104–105 greater than the concentration in the cytoplasm. In the case of the monoamines, this may correlate with actual difference between cytosolic and lumenal concentrations (Johnson, 1988; Parsons, 2000). Since an increase in acidification of the vesicle appears to increase transmitter content (Pothos et al., 2002), and a decrease in acidification decreases quantal size (Zhou et al., 2000; Pothos et al., 2002), strength of the electrochemical gradient may, in some cases, directly regulate the lumenal concentration of neurotransmitter. However, as noted above, other parameters are also likely to be important in determining transmitter transport and storage into secretory vesicles. Thus, the importance of DmH+ notwithstanding, it is also important to obtain more direct measurements of transmitter content. The potential to readily oxidize monoamine transmitters has been particularly useful in this regard. In carbon fiber amperometery, oxidation of individual amine molecules can be converted to a quantifiable electric current (Wightman et al., 1991). This technique has been used to directly measure the number of dopamine molecules that are released from an individual SV as 7,400+/700 (SEM) (Pothos et al., 2000). In organotypic cultures from rat brain, amperometry has been used to estimate the release of 31, 000 to 47,000 molecules of NE from a single SV (Chiti and Teschemacher, 2007). Both biochemical as well as electrophysiological assays (to compare the amount of transmitter that will give a post-synaptic response similar to that caused by a single SV) have been used to measure quantal size

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for ACh. Estimates for the number of acetylcholine molecules per SV are in the range of 7,000 (Kuffler and Yoshikami, 1975) to 12,000 (Miledi et al., 1983; Van der Kloot, 1990). Assuming an average content of 10,000 molecules of ACh, the concentration of a spherical vesicle, 40 nm in diameter would be 500 mM. The glutamate content of an individual SV appears to be somewhat lower than either amines or ACh, and in the order of 2,000 (Rao-Mirotznik et al., 1998) to 3,600 molecules (Riveros et al., 1986) per vesicle, at concentrations of 60 mM (Burger et al., 1989) to 210 mM (Riveros et al., 1986). These lower values relative to acetylcholine and amines may be the result of differences in either transport or storage and possibly higher glutamate efflux from synaptic vesicles (Carlson and Ueda, 1990). It should be noted, however, that serotonin as well as most other amines can also ‘‘leak’’ out of vesicles (Michalke et al., 1990; Floor et al., 1995; Romanenko et al., 1998). In addition to synaptic vesicles, large dense core vesicles (LDCVs) in neurons, and large dense core granules (LDCGs) in neuroendocrine cells can store monoamines (Liu et al., 1999; Torrealba and Carrasco, 2004) and perhaps acetylcholine (Lundberg et al., 1981; Agoston and Whittaker, 1989; Krantz et al., 2000). In addition to differences in size, localization and release properties, these vesicle types contain a dense proteinaceaous core, which is visible in electron micrographs (Torrealba and Carrasco, 2004). It has been suggested that lumenal proteins in SVs may bind neurotransmitter, but this has not been clearly established (Partilla et al., 2006). Transmitter that binds to the dense core of LDCVs and LDCGs is thought to be removed from free solution and thus from the constraints of osmolarity. This has been generally assumed to allow concentrations in LDCVs and LDCGs to be higher than those in SVs. However, amperometric measurements of LDCVs vary from 100 mM (Grabner et al., 2005), 300 mM (Wightman et al., 1991), 700 mM (Albillos et al., 1997) and up to 2.5 M (Ales et al., 1999). Furthermore, predicted concentrations of serotonin in SVs and LDCVs released from central neurons may be similar in some cases (Bruns et al., 2000). For dopamine, carbon fiber amperometry measurements show that the number of dopamine molecules per LDCV in PC12 cells is in the order of 73,000, versus 7400 for SVs derived from midbrain neurons (Pothos et al., 2000). Assuming a diameter of 80–100 nm, this yields an estimated concentration of 200 to 500 mM for dense core vesicles. Similarly, dense core granules in chromaffin granules are estimated to have lumenal concentrations of 500 mM based on histochemical labeling experiments (Hillarp, 1959), and 600 mM using biochemical assays (Phillips, 1982). In sum, it is possible that LDCGs and LDCVs do indeed contain more neurotransmitter than SVs, but this may not be the case for all neurons and neuroendocrine cells.

4.3 Variation in Quantal Size Both biochemical and electrophysiological studies have suggested that the numbers quoted above are not constant, and that quantal size the amount of neurotransmitter in a single vesicle can vary for both SVs and LDCVs, both at the neuromuscular junction (reviewed in (Van der Kloot, 1990, 2003)) and in the central nervous system (reviewed in (Edwards, 2007) (also see below, > Section 10.1) Potential mechanisms governing quantal size at the level of the vesicle itself have been divided into those dependent on the lumenal concentration of transmitter, the volume of the vesicle, or a set point that might operate independently of either variable (Edwards, 2007; Williams, 1997). Additional mechanisms based on rate at which the fusion pore opens are also possible, but seem less likely to depend on the function of vesicular transporters and are therefore beyond the scope of this chapter. Classical biochemical experiments using SVs derived from cholinergic tissues have established that at least two classes of SVs, termed VP1 and VP2, are present in cholinergic nerve terminals (Zimmerman and Whittaker, 1977). VP2 vesicles recycle more rapidly, are smaller, and contain less acetylcholine than VP1 vesicles (Prior and Tan, 1995). Moreover, the amount of vesamicol that binds to VP1 versus VP2 vesicles also varies, suggesting differences in the number of VAChT molecules per vesicle (Gracz et al., 1988). Similarly, EM analysis has shown that the Drosophila neuromuscular junction contains two distinct sizes of glutamatergic SVs. Parallel electrophysiological measurements show that the two vesicle populations generate different post-synaptic responses (Karunanithi et al., 2002). In leech serotonergic Retzius neurons, both amperometric methods and EM can distinguish different vesicle populations, and also show a

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correlation between quantal size and diameter of each population of vesicles (Bruns et al., 2000). In chromaffin cells, capacitance can be used to measure vesicle volume, and coupled with cell-attached patch amperometry, it has been used to show that the catecholamine content in dense core vesicles is also proportional to their volume (Gong et al., 2003). These studies suggest that transmitter concentration may remain constant while vesicle size changes. However, the debate over quantal size is not over and it is possible that several different mechanisms influence transmitter content depending on the preparation. Capacitance measurements at the glutamatergic calyx of Held do not appear to correlate with mini size (Wu et al., 2007). Furthermore, larger events were found to be less responsive to receptor blockade or dialysis. Together, this data was taken as an indication that, in this preparation, the concentration of neurotransmitter in the vesicle rather than vesicle volume is the major determinant of quantal size (Wu et al., 2007). Assuming for the moment that volume is the relevant parameter in most preparations, it is useful to consider the mechanism by which volume of the vesicles might change. On the one hand, an increased vesicle volume may result from osmotic equlibration of water content as the solute content of the vesicle lumen increases. However, unless additional lipid is added, this mechanism implies that the vesicle membrane can stretch. Conversely, it has been suggested that membrane may be added or subtracted to drive the increase in vesicle volume and quantal size. As discussed (Gong et al., 2003) this could be accomplished by cytosolic phospholipid transport proteins, or fusion of vesicles via membrane trafficking. Importantly the second mechanism could result in the addition of both lipids and integral membrane proteins. As we discuss in more detail below, the expression of vesicular transporters has a dramatic effect on quantal size, and vesicular transporter expression also increases vesicle volume. These observations suggest that the addition of membrane plus transporter(s) may serve to increase quantal size. This model remains speculative but would be consistent with at least some of the current data.

4.4 Rate of Transport When synaptic vesicles are rapidly recycled, the rate of vesicular transport may determine how full they will be at the moment of release. Turnover rate for human VAChT expressed in PC12 cells is 65 min1 (Varoqui and Erickson, 1996). Turnover rate for bovine VMAT in chromaffin granules has been reported to be 120 min1 (Scherman and Boschi, 1988) to140 min1 (Gasnier et al., 1987). Amine transport in homogenates derived from the CNS, presumably mediated by VMAT2, has been reported to be 10–35 min1 (Scherman, 1986), and heterologously expressed VMAT1 and 2 yield turnover rates of 10–40 min1 (Peter et al., 1994). It is useful to consider whether a rate similar to these would suffice to fill a vesicle in the time required for recycling by endocytosis. This has been estimated for cholinergic vesicles to be approximately one minute (Betz and Bewick, 1992), and in some synapses may be as fast as fifteen seconds (Ryan and Smith, 1995). Is this a sufficient time to fill a vesicle? At the frog NMJ, repetitive stimulation decreases quantal size and return to baseline is blocked by vesamicol (Naves and Van der Kloot, 2001). This data suggests that under conditions of rapid recycling and sustained release, VAChT turnover rate may not be able to keep up with the rate of SV recycling, or keep a recycling vesicle filled with ACh. Conversely, repetitive stimulation of cultured glutamatergic neurons does not decrease quantal size (Zhou et al., 2000). It is possible that the turnover rate for VGLUT is exceptionally high, or the number of VGLUTs per SV exceeds the number of VAChT proteins per SV. If we estimate the content of glutamate in an SV at 3,000 molecules per SV (see above), and the turnover number of VGLUT to be 300 min1 (approximately double that of native VAChT), one VGLUT could fill a vesicle in 10 min and ten VGLUTs could do it within 1 min. We return to transporter stoichiometry below, and note that estimates vary from 1 to 10 per vesicle.

4.5 When are Vesicles Loaded? It is worth noting that other parameters affect packaging of neurotransmitter into SVs. Early biochemical studies of acetylcholine and amine transport have shown that recently synthesized transmitter may be

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preferentially released (Besson et al., 1969; Molenaar et al., 1971). In addition, vesamicol preferentially releases recently transported acetylcholine out of the vesicle lumen (Anderson et al., 1986). The idea that additional acetylcholine may be added to vesicles immediately prior to release has been called ‘‘second stage loading,’’ and under some conditions, may be responsible for up to 30% of the transmitter released from a single SV at the frog NMJ (Doherty et al., 1984; Naves and Van der Kloot, 1996). SVs that transport newly synthesized acetylcholine may be preferentially released, and also show higher vesicular acetylcholine transport activity (Gracz et al., 1988). It is possible that these effects involve alterations in the function of VAChT and the VMATs (Gracz et al., 1988).

4.6 Transporter Substrates and Their Affinities Substrate affinity varies considerably between different vesicular transporters. Since the transport cycle requires binding and unbinding of substrate from at least two distinct conformations, at least two substrate affinity constants (Kd) are involved in transport. The actual Kds for binding of substrate to vesicular transporters can be measured directly, and rate constants have been derived for each step in the transport cycle (Parsons, 2000). However, in most studies, Michaelis–Menton parameters are used to represent activity of the entire transport cycle for ACh. This has also been referred to as the apparent or macroscopic Km. IC50 measurements, rather than apparent Kms which have been obtained in many studies.We therefore refer below more generally to the apparent affinity of the transporters when discussing Km and IC50 values. For VAChT and VMATs, apparent Kms derived from transport assays are 10–100 fold smaller than Kd values; this difference is thought to be caused by kinetic contributions to apparent affinity (Parsons, 2000). Both cell free (Peter et al., 1994) and whole cell assays (Erickson et al., 1996) have been used to analyze the relative apparent affinity of VMAT1 and VMAT2 for monoamine substrates in heterologously transfected cells. Relative affinities across different amines are similar for both assays, with 5HT>DA, Epi, NE>>histamine1. However, the absolute values are generally lower for cell free versus whole cell assays with VMAT2 showing apparent affinities for 5HT, DA, Epi, NE, histamine of 0.19, 0.32, 0.47, 0.33, 3.06 mM (cell free) (Peter et al., 1994) versus 0.9, 1.4, 1.9, 3.4, 143 mM (whole cell) respectively (Erickson et al., 1996). For VMAT1, apparent affinities (micromolar) for 5HT, DA, Epi, NE, histamine are 0.85, 1.56, 1.86, 2.5, 436 (cell free) as against 1.4, 3.8, 5.5, 13.7, 4,696 (whole cell) respectively. Thus, in general, apparent affinity of VMAT2 for most monoamines is greater than VMAT1; one known exception is tryptamine, which has a higher affinity for VMAT1 (IC50 of 0.4 mM versus 2.19 mM for VMAT2) (Finn and Edwards, 1997). Interestingly, the apparent affinity of histamine for both transporters is dramatically lower than for other amines, and histamine is not likely to be a physiological substrate for VMAT1, since only VMAT2 is expressed in histaminergic cells (see below). Residues responsible for the affinity of VMAT2 for histamine include multiple sites in the 12 transmembrane (TM) ‘‘backbone’’ (Peter et al., 1996; Finn and Edwards, 1997, 1998) (see also > Section 6.3). The role of the trace amines tyramine and octopamine in the mammalian nervous system remains unclear. In contrast, octopamine clearly functions as a neurotransmitter in invertebrates, and in arthropods, octopamine is thought to play a prominent role at the neuromuscular junction (Breen and Atwood, 1983; Rodriguez-Valentin et al., 2006). The Drosophila homolog of VMAT is expressed in octopaminergic nerve terminals in the fly, and DVMAT shows an IC50 for octopamine of 1 mM, as compared to an IC50 of 0.5 mM for dopamine using either Drosophila or mammalian VMAT in the same preparation (Greer et al., 2005). In addition to established endogenous substrates, VMATs recognize other charged molecules that do not function as neurotransmitters. These include the neurotoxin MPP+ as well as ethidium (Liu et al., 1992b; Yelin and Schuldiner, 1995). It is possible that other molecules are transported by VMATs into secretory vesicles in vivo, and that sequestration of toxins by VMAT plays a cytoprotective role (Yelin and Schuldiner, 1995) similar to eukaryotic multidrug transporters and bacterial TEXANs. VAChT may transport a number of structurally divergent organic molecules which are similar to acetylcholine only in that they carry a charge of +1 (Bravo et al., 2005). VAChT may also transport choline

1

5HT, Serotonin; Epi, Epinephrine; NE, Norepinephrine. See also List of Abbreviations and Acronyms.

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(Bravo et al., 2004a) albeit with a sevenfold lower apparent affinity than ACh. Despite these additional potential substrates, it is thought that only acetylcholine functions as an endogenous substrate for VAChT (Parsons, 2000). Thus, in contrast to VMATs, VAChT as well as VGAT and VGLUTs generally show a more restricted range of endogenous neurotransmitter substrates. In addition, VAChT, VGAT and VGLUTs differ from VMATs in that they show a relatively low apparent affinity for substrate. The apparent Km of rat VAChT for acetylcholine is 1 mM, three orders of magnitude lower than the apparent Km of VMAT for most amines. Substrates for VGAT include GABA and glycine and possibly the structurally related molecule beta–alanine (Christensen et al., 1991). The apparent affinities of VGAT for GABA and glycine are 5 and 25 mM, respectively (Gasnier, 2004). Because of low apparent affinity of VGAT/VIAAT for glycine it has been difficult to assay uptake in vitro. Disruption of GABA and glycine release in VGAT/VIAAT knockout mice indicates that both transmitters are substrates of VGAT/VIAAT in vivo (Wojcik et al., 2006). The only known endogenous substrate for the VGLUTs is glutamate. Notably, none of the three identified VGLUTs appear to transport the related amino acid aspartate (Naito and Ueda, 1985; Bellocchio et al., 2000; Takamori et al., 2000; Gras et al., 2002; Varoqui et al., 2002), leaving open the possibility that a novel aspartate transporter will be identified. Similar to VGAT and VAChT, the apparent affinity of VGLUTs for substrate is relatively low and in the mM range (Liguz-Lecznar and Skangiel-Kramska, 2007), perhaps to prevent depletion of glutamate from the cytoplasm.

5

Pharmacology

A number of pharmacologic agents which alter the function of VMATs, VAChT and VGLUT, but not of VGAT, have been characterized. In most cases, drugs and environmental agents which interact with the transporters inhibit transport activity. However, the pharmacology of the VMATs is more complex than that of other vesicular transporters, since it includes additional agents that promote efflux (see below). The most commonly used and extensively characterized VMAT inhibitors are reserpine and tetrabenazine - neither of which undergoes transport by VMAT. Reserpine and tetrabenazine differ in several important respects, including their proposed mechanism of action, putative binding sites, and relative affinity. Reserpine binds with high affinity to both VMAT1 and 2 (Peter et al., 1994) at, or very near, a substrate-binding site in VMATs on the cytoplasmic side of the transporter (Weaver and Deupree, 1982; Scherman and Henry, 1984; Rudnick et al., 1990). Interestingly, the affinity of reserpine for VMATs depends on the conformation of the transporter, and this changes in response to activation of the transport cycle by the proton gradient (Weaver and Deupree, 1982; Scherman and Henry, 1984; Darchen et al., 1989; Rudnick et al., 1990). In the absence of a proton gradient, the Kd for VMAT1 is 25–340 nM, but this changes to 30 pM in its presence (Parsons, 2000). Since reserpine and amines appear to bind to the same site on VMAT, these observations suggest that the amine-binding site is significantly altered by the orientation of the transporter relative to the membrane. The structural basis of these changes remain unclear, but it is attractive to consider the possibility that the affinity of the substrate binding site might be lower when VMAT is poised to release transmitter into the vesicle lumen. In contrast to reserpine, tetrabenazine interacts with VMAT at a site distinct from the substrate binding site, and activation of the proton gradient does not affect tetrabenazine binding (Peter et al., 1994; Scherman and Henry, 1984). In addition, unlike reserpine, the affinity of tetrabenazine for VMAT1 and 2 differs, and the potency of tetrabenazine to inhibit VMAT2 is tenfold higher than for VMAT1 (IC50s of 0.3 and 3 mM, respectively) (Peter et al., 1994). Exploiting this difference, chimeras of VMAT1 and VMAT2 have been used to map domains required for tetrabenazine binding (Peter et al., 1996). Additional site directed mutagenesis as well as photoaffinity labeling indicate that residues in transmembrane domains (TMs) 10–12 are particularly important for tetrabenazine binding to VMAT2 (Finn and Edwards, 1997, 1998; Sievert and Ruoho, 1997; Thiriot and Ruoho, 2001) (see also > Section 6 below). Interestingly, despite the apparent difference in binding sites, tetrabenazine inhibits reserpine binding, presumably via an allosteric mechanism (Darchen et al., 1989). In contrast to reserpine and tetrabenazine, some drugs which interact with VMATs promote efflux of transmitter out of the vesicle lumen rather than inhibiting transport. Most, if not all of these are VMAT

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substrates, and many are derivatives of amphetamines, including methamphetamine and 3,4 methylenedioxymethamphetamine (MDMA, also known as Ecstasy) (Fleckenstein et al., 2007). Amphetamine derivatives are also substrates of plasma membrane monoamine transporters, and these drugs promote efflux out of the cell as well as the vesicle lumen (Fleckenstein et al., 2007). Indeed, the specificity for the action of amphetamines for particular aminergic cells is likely to depend on their affinity for particular plasma membrane monoamine transporters, since most drugs appear to function equivalently to drive efflux out of synaptic vesicles (Fleckenstein et al., 2007). The precise manner in which amphetamines and their derivatives promote efflux from the vesicle lumen remains controversial: efflux out of the synaptic vesicles has been proposed to involve either direct exchange with substrate (Partilla et al., 2006) or neutralization of the proton gradient as a weak base, and subsequent diffusion of substrate into the cytoplasm (Sulzer and Rayport, 1990). A number of drugs in addition to reserpine, tetrabenazine and amphetamines have been shown to interact with VMATs. Although cocaine is primarily used as an inhibitor of the plasma membrane dopamine transporter, it also interacts with VMAT, albeit with much lower apparent affinity (0.35 mM for DAT versus 137 mM for VMAT2) (Reith et al., 1994). Two other DAT inhibitors, GBR 12909 and 12935, also inhibit dopamine uptake into brain synaptic vesicles (Reith et al., 1994). Furthermore, unlike cocaine, the concentrations of GBR 12909 and 12935 required for inhibiting VMAT2 (34–45 nM) appear to be relatively similar to those required for inhibiting DAT (1–6 nM). Additional VMAT inhibitors include ketanserin (Darchen et al., 1988), amiodarone (Haikerwal et al., 1999), and lobeline, which also interacts with plasma membrane monoamine transporters, as well as a series of synthetic compounds structurally similar to lobeline (Harrod et al., 2001; Zheng et al., 2005). Interest in the development of these novel VMAT inhibitors stems in part from their potential use in the treatment of psychostimulant abuse (Zheng et al., 2006). It should also be noted that tetrabenazine is sometimes used to treat movement disorders (Paleacu et al., 2004). In addition to drugs, several environmental toxins and pesticides have been shown to inhibit VMAT activity. These include the organochlorine pesticide hepatachlor and the structurally related, polychlorinated biphenyls (PCBs) (Mariussen et al., 1999; Miller et al., 1999; Mariussen and Fonnum, 2001; Greer et al., 2005). The inhibition of VMATs by heptachlor and PCBs may be relevant to Parkinson’s disease, since both PCBs as well as organochlorine pesticides have been shown to deplete dopamine stores in experimental animals (Kirby et al., 2001), and organochlorines may be associated with an increased incidence of Parkinson’s disease (Fleming et al., 1994; Dick, 2006). Vesamicol (L-trans-2-(4-phenyl[3,4-3H]piperidino)cyclohexanol) is the major pharmacologic agent used in the study of VAChT (Bahr and Parsons, 1986; Parsons, 2000). Prior to the molecular identification of VAChT, the vesicular protein known to be involved in acetylcholine storage was sometimes called the vesamicol receptor, and vesamicol can be used to affinity-purify VAChT (Bahr and Parsons, 1992; Bahr et al., 1992a). Additional structural derivatives of vesamicol have been synthesized as alternative probes for VAChT (Wenzel et al., 2005). Vesamicol binds to the cytosolic side of VAChT (Noremberg and Parsons, 1989) with an equilibrium dissociation constant Kd of 5 nM (Bahr and Parsons, 1986). Although binding sites for vesamicol and acetylcholine are distinct, they are thought to be close to one another thus allowing some functional interactions (Bahr et al., 1992a; Kim et al., 2000; Parsons, 2000; Zhu et al., 2001; Ojeda et al., 2004; Bravo et al., 2004b), for example vesamicol competes with acetylcholine for binding to VAChT when the substrate binding site is oriented toward the cytoplasm (Bravo et al., 2004b). The pharmacology of VGAT is poorly defined, and to our knowledge, specific inhibitors of VGATs have not yet been developed. Gamma-hydroxybutyrate (GHB), notorious as a sedative-hypnotic, has been suggested to be substrate of VGAT (Muller et al., 2002). Recognition and storage of GHB by VGAT could conceivably be involved in the putative actions of this drug at the synapse (Muller et al., 2002). Multiple vesicular glutamate transport inhibitors have been characterized, although inhibitors specific for particular VGLUT isoforms have not been identified. Commonly used VGLUT inhibitors include azo dyes such as Evans Blue and Trypan Blue, which show a Kis of 87 and 49 nM, respectively (Roseth et al., 1998). Other VGLUT inhibitors include Rose Bengal (Ogita et al., 2001), bromocryptine - better known for its effects on dopaminergic signaling (Carlson et al., 1989b) - and the anion channel inhibitor 4,4-diisothicyanatostilben- 2,2 – disulfonic acid (DIDS) (Hartinger and Jahn, 1993). In addition, a series

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of quinoline dicarboxylate derivatives have been synthesized and tested as VGLUT inhibitors (Fonnum et al., 1998; Roseth et al., 1998; Carrigan et al., 1999, 2002). Although no current VGLUT inhibitors are used clinically, their development would be potentially useful, since a non-toxic VGLUT inhibitor that could easily enter the CNS might be used to combat the cytotoxic effects of excessive glutamatergic signaling (Reimer and Edwards, 2004).

6

Models and Structure/Function Studies

6.1 Models To date, an X-ray crystallographic structure is not available for any of the vesicular transporters. However, structural and functional information for related members of the MFS have been used to help generate working models of vesicular transport. Crystal structures are now available for Lac permease (Abramson et al., 2003; Guan et al., 2007), the glycerol-3-phosphate transporter (Huang et al., 2003), the oxalate transporter (Hirai et al., 2002), and the bacterial multidrug transporter (Yin et al., 2006). Together with the extensive biochemical characterization of Lac permease, and the biochemical and molecular characterization of VAChT and VMAT, structures of related MFS members have been used to create a working model of acetylcholine transport (Bahr et al., 1992a; Parsons, 2000; Bravo et al., 2004b). In the current working model of vesicular acetylcholine (and possibly amine) transport, the exchange of one lumenal proton is used to transport substrate into the vesicle lumen, and a second proton is used to reorient the substrate binding site back to the cytoplasm to allow the cycle to begin again (Bahr et al., 1992a; Parsons, 2000). Acetylcholine binding is likely to be at equilibrium; in contrast, the two proton-dependent movements of the transporter are thought to be rate limiting, and reorientation of the empty acetylcholine binding site is likely to be the slower of the two translocation events (Bravo et al., 2004b). Whether or not each element of this model is correct, it continues to provide a crucial framework for many of the structure– function experiments described below.

6.2 Membrane Topology For VMATS and VACHT, most but not all computer-based predictions suggest 12 TMs with the N and C termini facing the intracellular milieu (Eiden et al., 2004) and a larger lumenal loop between the first and second transmembrane domains. Biochemical studies support these predictions insofar as the C termini of both VAChT and VMAT2 are phosphorylated and interact with the cytosolic trafficking machinery (Krantz et al., 1997, 2000; Cho et al., 2000; Kim and Hersh, 2004). Similarly, the large loop between TMs 1 and 2 is glycosylated and epitope tags in this domain are exposed to the extracellular milieu, supporting its predicted localization to the vesicle lumen (Tan et al., 1998; Cho et al., 2000). Topological predictions for VGLUTs have been less consistent, and range from 10 to 12 TMs. Biochemical data including the association of the N- and C-termini with cytosolic reagents confirm that both termini face the cytoplasm (De Gois et al., 2006; Jung et al., 2006; Voglmaier et al., 2006). The topology of the rest of the protein has been investigated using epitope tags placed in predicted cytoplasmic and lumenal domains (Fei et al., 2007). The data Suggest ten bona fide TMs with two additional regions that are either contained in the bilayer or minimally exposed to the vesicle lumen for a total of 12 membrane-bound domains (Fei et al., 2007). Topology of VGAT has not been tested experimentally but is predicted to include ten TMs (McIntire et al., 1997; Sagne et al., 1997).

6.3 Structure/Function Studies of Transport Activity The ‘‘backbone’’ of VAChT and VMAT may be defined as the twelve TMs and intervening loops, exclusive of the N- and C-termini. The backbone of VMAT and VAChT includes most if not all the residues required for transport activity and substrate recognition (Fisher et al., 1993; Peter et al., 1996; Finn and Edwards, 1997, 1998; Kim et al., 1999; Thiriot and Ruoho, 2001; Thiriot et al., 2002; Ojeda et al., 2004) (> Figure 7-1a, b).

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. Figure 7-1 Membrane bound backbone of the vesicular transporters mediates transport activity. Vesicular transporters are polytopic integral membrane proteins: VMAT2 (a) and VAChT (b) are likely to contain 12 transmembrane domains. VGLUT1 (c) also contains 12 membrane-bound domains, but only ten appear to completely span the bilayer. (VGAT is predicted to contain ten transmembrane (TM) domains but this has not been experimentally tested and is not shown). Site directed mutagenesis has revealed the function of several residues in (a) VMAT2, (b) VAChT and (c) VGLUT1. For VMAT2 these include aspartates (D) in the first and tenth membrane-bound domains required for transport activity, a charge-pair between residues in TM2 (K) and TM11 (D), and a proposed disulfide bond between cysteines (C) in two lumenal loops. Residues important for substrate affinity include a tyrosine (Y) in TM11 and an aspartate (D) in TM12. For VAChT, a tryptophan (W) in TM8 is required for affinity to ACh. Conserved regions that contain glycine and/or proline (G/P) and an aspartate in TM11 (D) are required for transport activity. For VGLUT1, mutation of the arginine (R), histidine (H) and glutamate (E) residues in the indicated membrane-bound domains have been shown to decrease transport activity. Note that location of residues in the cartoon is approximate and does not accurately reflect their position within each domain

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These include several charged residues embedded in predicted transmembrane domains: For both rat VMAT2 and VAChT, mutation of an aspartate in the tenth transmembrane domain (TM10) abolishes transport (Merickel et al., 1997; Kim et al., 1999); glutamate can substitute for aspartate, indicating the importance of an acidic residue at this site (Merickel et al., 1997; Kim et al., 1999). Mutation of an aspartate in TM1 of VMAT2 but not VAChT also inhibits transport (Merickel et al., 1997; Kim et al., 1999). Additional charged residues in TM2 and TM11 of VMAT2 are thought to form a charge pair (Merickel et al., 1997) (> Figure 7-1a). The charges can be exchanged without disrupting transport, underscoring the likelihood that they interact, and that TM2 and TM11 may be in direct contact. However, the analogous aspartate in TM11 of VAChT does tolerate exchange with its proposed basic partner, suggesting that either VAChT has a more stringent structural requirement, or that this residue is more directly involved in substrate translocation (Kim et al., 1999). Additional residues that are likely to have structural importance in VMATs include cysteines in the large lumenal loop between TMs 1 and 2 and the loop between TMs 7 and 8; these two residues are thought to form a disulfide bond (Thiriot et al., 2002) (> Figure 7-1a). VMAT1 and 2 diverge in both substrate and drug affinity but are structurally similar and therefore likely to use the same transport mechanism. Chimeric constructs fusing VMAT1 and 2 show active transport (Peter et al., 1996), and have been used to map domains important to substrate and drug affinity. In general, TMs 5–8 and 9–12 show the greatest contribution, and additional experiments using point mutants in these regions have identified the importance of several specific amino acids (Finn and Edwards, 1997, 1998). A tyrosine in TM11 (Y434 in VMAT2) and an aspartate in TM12 (D461) are particularly important for higher affinity of VMAT2 for serotonin and possibly histamine, as well the drug tetrabenazine (Finn and Edwards, 1997) (> Figure 7-1a). The apparent affinity of dopamine was not affected by mutation of Y434 suggesting that the ring nitrogen present in serotonin and histamine, but not dopamine, may interact with this residue (Finn and Edwards, 1997). Y434 in VMAT2 is also responsible for the relatively low affinity of VMAT2 for tryptamine, and underscores the possibility that this residue directly interacts with substrate (Finn and Edwards, 1997). A similar analysis of residues TMs 5–8 in the VMATs showed that mutations in this region could reverse the effects of mutations in TM9–12 (Finn and Edwards, 1998), suggesting that the two regions may interact. Site directed mutagenesis studies on VAChT have taken advantage of crystal structures available for other proteins that bind ACh. For these proteins and for VAChT, it has been hypothesized that pi electrons in the ring of aromatic residues may interact with the positive charge in acetylcholine (Ojeda et al., 2004). Mutation of multiple candidates has highlighted the importance of one aromatic residue, a tryptophan in TM8 at position 331 (W331), for which substitution decreased acetylcholine affinity but did not otherwise interfere with the function of the transporter (> Figure 7-1b). This data supports the idea that pi electron solvation of W331 could potentially be involved in binding acetylcholine to VAChT. More fundamentally, they suggest this site may help form the substrate-binding pocket in VAChT (Ojeda et al., 2004). Some studies have attempted to map the site of proton translocation in VAChT and VMAT, based in part on the idea that conserved histidines may play an important role (Kim et al., 2000; Parsons, 2000; Shirvan et al., 1994). Although some possibilities have been suggested (Shirvan et al., 1994; Kim et al., 2000), the identity of the residues that accept and release protons during the transport cycle remain unresolved (Parsons, 2000). Additional studies have identified amino acids in VAChT that may be involved in conformational shifts thought to be required for translocation of substrate. In other members of the MFS, including Lac permease, it has been suggested that the substrate may be transported through a channel-like domain, with conformational changes which rock the opposing sides of the channel allowing transport (Abramson et al., 2004; Hirai and Subramaniam, 2004; Lemieux et al., 2004; Kaback et al., 2007). Because the structures of the crystallized MFS transporters are similar, it has been argued that conserved motifs may be responsible for the structural shifts that allow transport (Hirai et al., 2003; Vardy et al., 2004; Chandrasekaran et al., 2006). In support of this idea, the mutation of residues contained within conserved proline- and glycinerich regions in VAChT block transport activity (Chandrasekaran et al., 2006). In contrast to VMAT and VAChT, relatively few structure-function experiments have been performed on the presumptive VGLUT and VGAT transport domains. In the case of VGLUTS, several TMs contain potentially charged residues (TM 1, 2, 4, 7). Some of these are highly conserved among members of the

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extended SLC17 family including sialin and the NaPi transporters (Juge et al., 2006). In the one study published to date, it was shown that mutation of Arg184 (TM4), His128 (TM2), and Glu191 (TM4) dramatically decreased VGLUT transport activity (Juge et al., 2006) (> Figure 7-1c). Based on modeling with the bacterial glycerol 3 phosphate transporter, all three cluster together in a domain predicted to bind substrate (Juge et al., 2006). To our knowledge, nothing has been published on the amino acids in VGAT that are required for transport.

7

Expression

7.1 The CNS Vesicular neurotransmitter transporters are expressed both in the CNS and the periphery. We review first their expression in the CNS. Although mammals express two VMATs, only VMAT2 is expressed in the mature central nervous system (Peter et al., 1995; Erickson et al., 1996). VMAT2 is expressed in dopaminergic, serotonergic, and adrenergic cells in the brainstem and all of their abundant projections; thus, VMAT2 is thought to be responsible for storage of all monoamines in the CNS (Peter et al., 1995; Erickson et al., 1996). VAChT has been shown to be co-expressed with ChAT in all the major, previously identified cholinergic cells in the nervous system. These include clustered cell bodies in known cholinergic nuclei (e.g., in the basal forebrain) and the more widely dispersed interneurons of the striatum (Gilmor et al., 1996; Weihe et al., 1996). In addition, the immunohistochemical and mRNA in situ analysis of VAChT has allowed the identification of additional cholinergic projections difficult to visualize using other markers; these include a dense plexus of cholinergic projection in the median eminence and cell bodies in the hypothalamus that may be the source of these processes (Weihe and Eiden, 2000). Only one isoform of VGAT/VIAAT has been identified and it is expressed in GABAergic and glycinergic neurons throughout the CNS (Chaudhry et al., 1998). In the eye, this includes the horizontal and amacrine cells, and some bipolar cell in the retina (Cueva et al., 2002; Jellali et al., 2002). VGLUT1 and 2 show essentially complementary patterns of expression in the CNS (> Figure 7-2). VGLUT1 is the primary isoform expressed in the telencephalon, including the cortex and hippocampus . Figure 7-2 VGLUT 1 and 2 have complementary patterns of distribution (see text for details). The cartoon shows structures in adult animals in which either VGLUT1 or VGLUT2 predominates. It should be noted that during development VGLUT 1 and 2 are co-expressed in some structures such as the cerebellum

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(Boulland et al., 2004; Fremeau et al., 2004b). VGLUT2 is the primary isoform in the diencephalon and rhomencephalon, including the pons, thalamus, hypothalamus, and the spinal cord (Varoqui et al., 2002; Boulland et al., 2004; Liguz-Lecznar and Skangiel-Kramska, 2007). These patterns cover many of the known glutamatergic cells in the brain (Varoqui et al., 2002; Boulland et al., 2004; Fremeau et al., 2004b). It should be emphasized, however that although one isoform may predominate, both VGLUT1 and 2 are expressed to some extent in many structures (Liguz-Lecznar and Skangiel-Kramska, 2007). Electrophysiological assays also suggest that VGLUT1 and 2 localize to relatively distinct sites in the mature CNS. Slice preparations from VGLUT1 knockout mice show a dramatic, overall decrease in glutamatergic signaling, but a number of glutamatergic synapses remain active, presumably as a result of glutamate release by VGLUT2 (Fremeau et al., 2004b). Additional experiments using the drug MK801 to block post-synaptic glutamate receptors in an activity-dependent fashion show wild type rates of inactivation in the VGLUT1 knockout mice, suggesting that at the remaining active synapses, glutamate release is not reduced. This data has been interpreted as indicating that VGLUT 1 and 2 release glutamate at different, mutually exclusive sites, at least in the mature nervous system (Fremeau et al., 2004b). In contrast, it is likely that VGLUT1 and 2 co-localize during development (see > Section 7.5). VGLUT3 is expressed in scattered loci throughout the CNS, including regions in the telencephalon and diencepahlon that are otherwise dominated by VGLUT1 and 2 (Fremeau et al., 2002; Gras et al., 2002; Schafer et al., 2002). Similar to VGLUTs 1 and 2, VGLUT3 may be expressed in some glutamatergic cells, possibly including pyramidal cells of the neocortex (Schafer et al., 2002; Fremeau et al., 2002; Gras et al., 2002; Harkany et al., 2004). However, in striking contrast to VGLUTs 1 and 2, VGLUT3 is primarily expressed in cells that have not been classified as glutamatergic. Therefore, as we discuss in more detail below, VGLUT3 is co-expressed with other vesicular transporters (Fremeau et al., 2002; Gras et al., 2002; Schafer et al., 2002).

7.2 Glia Classically, exocytosis in the nervous system has been thought to be confined to neurons. More, recent data suggests that glutamate is also released from glia (Bezzi et al., 2004; Montana et al., 2006). Consistent with this idea, both VGLUT1 and 2 as well as VGLUT3 have been shown to be expressed in astrocytes and possibly oligodendrocytes (Boulland et al., 2004; Montana et al., 2004). Glutamate release from glia has been shown to signal neurons via glutamate receptors at apposed, non-synaptic sites (Jourdain et al., 2007). This in turn may regulate neuronal glutamate release at the synapse (Jourdain et al., 2007).

7.3 Mismatched Expression Our discussion of transporter expression patterns in the CNS would not be complete without mentioning the occasional mismatch in expression observed for VMAT2 and the enzymes responsible for monoamine biosynthesis. Most catecholaminergic neurons express both VMAT2 as well as tyrosine hydroxylase (TH) and amino acid decarboxylase (AADC, also known as Dopa decarboxylase). However, an additional subset of aminergic cells that express TH and AADC do not express detectable levels of VMAT2 (Weihe et al., 2006). These include aminergic neurons in the olfactory bulb and nucleus tractus solitarius (Weihe et al., 2006). It remains unclear whether dopamine is released from these cells. An additional anomaly is the expression of VMAT2 in some cells that do not synthesize monoamines. In the rodent brain, several neuronal subtypes that are glutamatergic in the adult express VMAT2 and the plasma membrane serotonin transporter (SERT) during development (Lebrand et al., 1996, 1998). This phenomenon has been studied extensively for thalamocortical neurons that project to the barrel cortex of rodents and form a pattern representing input from specific whiskers (whisker barrels). Since thalamocortical projections do not express the enzymes required for serotonin biosynthesis, it has been proposed that these cells take up serotonin from the extracellular milieu rather than synthesizing it themselves (Lebrand et al., 1996). A similar phenomenon has been suggested to occur in the somatosensory cortex of primates

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(Lebrand et al., 2006). Pharmacologic and genetic disruption of Monoamine Oxidase (MAO) and/or SERT in rodents causes defects in the barrel fields, indicating that amine release has an important developmental role (Alvarez et al., 2002; Cases et al., 1996). The mechanism by which serotonin affects patterning in the cortex may include serotoneric regulation of the axon guidance molecule netrin-1 (Bonnin et al., 2007). VMAT2 knockout mice do not show as severe a disruption in barrel formation as MA0-A knockout mice, suggesting that non-exocytotic release of serotonin may be important for some aspects of cortical development (Persico et al., 2001). However, the normal pattern of at least one subset of neurons is disrupted in VMAT2 knock out mice, consistent with a functional role for VMAT2-mediated, exocytotic release of serotonin in some aspects of thalamocortical projections during development (Alvarez et al., 2002).

7.4 Outside the CNS Neurons in the periphery as well as the CNS express vesicular transporters. As in the CNS, peripheral cells that release monoamines usually express only one VMAT isoform: either VMAT1 or VMAT2 (however, see > Section 7.6 below). Enterochromaffin cells in the small and large intestine primarily express VMAT1 for the storage of serotonin (Peter et al., 1995; Erickson et al., 1996; Weihe and Eiden, 2000). These include serotonergic cells in the myenteric plexus (Weihe and Eiden, 2000). VMAT1 is also expressed in pinealocytes where it has been proposed to store serotonin (Hayashi et al., 1999) and in dopaminergic cells in the kidney proximal tubule (Maurel et al., 2007). VMAT2 is expressed in most adrenergic neurons of the sympathetic ganglia where it allows storage of norepinephrine. It is also expressed in enterochromaffin-like cells of the gastric mucosa, which release histamine in response to gastrin, in mast cells for storage of both histamine and serotonin (Peter et al., 1995; Erickson et al., 1996; Travis et al., 2000) and in platelets, presumably for storage of serotonin (Anlauf et al., 2006). Although it is difficult to easily categorize a universal pattern for the expression of VMAT1 and VMAT2 in the periphery, one important generalization is that VMAT2 is expressed in cells that store histamine, consistent with its higher apparent affinity for this transmitter. Additional peripheral sites of vesicular transporter have been reported, but the function of neurotransmitter release at these sites remains poorly understood. VGLUT3 is expressed in the liver (Fremeau et al., 2002) and both VGLUT1 and 2 are expressed in the pancreas; VGLUT2 is also expressed in the stomach, intestine, and testes (Hayashi et al., 2003). Beta cells of the pancreas express both VMAT2 (Anlauf et al., 2003) and VGAT (Mayerhofer et al., 2001). Interestingly, beta cells may release GABA and serotonin from the same secretory vesicles (Braun et al., 2007) suggesting that VGAT and VMAT2 may co-localize to the same secretory vesicles in these cells (Takahashi et al., 1997a) (see also > Section 7.6 below).

7.5 Developmental Changes Developmental studies of transporter expression have been performed for VMATs (Weihe and Eiden, 2000), VGLUTs (Boulland et al., 2004) and VGAT (Oh et al., 2005). A developmental change in expression from VMAT1 to VMAT2 occurs in the dopaminergic glomus cells of the carotid body (Weihe and Eiden, 2000). These cells function as sensors for oxygen and thus help to regulate respiration. In the adult, they express VMAT1, but early in development they also express high levels of VMAT2 (Weihe and Eiden, 2000). Transient expression of VMAT1 mRNA also has been reported in the CNS (Hansson et al., 1998); however, subsequent studies have failed to reproduce this finding (Weihe and Eiden, 2000). Absence of detectable monoamine storage in the CNS of VMAT2 knockout mice supports the notion that only VMAT2 is expressed in the CNS in rodents (Fon et al., 1997). VGLUT1 and 2 also show developmental changes in expression. VGLUT2 shows high levels of expression after birth that later decline in a number of regions: this pattern is most prominent in the cerebellum (Boulland et al., 2004). By adulthood, VGLUT2 is essentially absent from the cerebellum and has been replaced by VGLUT1 (Boulland et al., 2004). In a few cases, VGLUT1 and 2 have been shown to transiently co-localize in the same cell, e.g., in somatosensory cortex and ventral posteromedial thalamic nucleus (Nakamura et al., 2005).

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From a functional standpoint, the reason why the expression of VGLUT1 and 2 might change during development is not yet clear. The kinetics of glutamate transport VGLUT1 and 2 do not differ dramatically, suggesting that neurotransmitter loading into SVs would not be affected by switching from one VGLUT isoform to another. However, VGLUT1 and 2 contain different trafficking motifs (Voglmaier et al., 2006) (see > Section 8.2) and changing from VGLUT2 to 1 may reflect differential developmental requirements for SV trafficking, e.g., the use of an alternate mode to recycle transporters to SVs early in development.

7.6 Co-Expression Some cells in the adult CNS may express two different types of vesicular transporters, consistent with the relatively recent idea that two different classical neurotransmitters can be stored and released from the same cell (Trudeau, 2004; Seal and Edwards, 2006). During a transient peak in VGLUT3 expression at postnatal day 10 in the rat, VGLUT3 co-localizes with VGAT in the soma and terminals of Purkinje cells (Gras et al., 2005), potentially allowing the transient release of GABA and glutamate. In the primate autonomic nervous system, VAChT and VMAT2 are co-expressed during development in several parasympathetic ganglia (Schutz et al., 1998; Weihe et al., 2005). However, since these cells do not express the biosynthetic enzymes for catecholamine synthesis, it is unclear whether VMAT2 is used here for transmitter release. Rather, it is possible that VMAT2 may serve another role, such as neuroprotection in these cells. The most surprising examples of co-expression include instances in the adult animal that are likely to result in the co-release of two different neurotransmitters from mature neurons. The co-localization of VGLUT3 in non-glutamatergic cells is the most dramatic example of this phenomenon (Fremeau et al., 2002; Gras et al., 2002; Schafer et al., 2002; Boulland et al., 2004). VGLUT3 is co-expressed with VAChT in most, and possibly all cholinergic interneurons in the striatum (Fremeau et al., 2002; Gras et al., 2002; Schafer et al., 2002). Co-localization with cholinergic markers in other regions such as the septum and basal forebrain are less prominent (Fremeau et al., 2002; Schafer et al., 2002); nonetheless a small subset of cholinergic cells that project to the basolateral amygdaloid nucleus also express VGLUT3 (Nickerson Poulin et al., 2006). GABAergic neurons cells in the hippocampus and neocortex co-express VGLUT3, including GABAergic interneurons that innervate pyramidal cells in the neocortex (Fremeau et al., 2002). VGLUT3 is also co-expressed with VMAT2 in a relatively large number of serotonergic processes that emanate from the raphe nucleus (Gras et al., 2002; Schafer et al., 2002; Boulland et al., 2004). Both VGLUT1 and 2 also appear to be expressed in neurons that release other transmitters, but in a somewhat more limited pattern than VGLUT3. A subset of bipolar cells in the retina expresses both VGAT and VGLUT1 (Kao et al., 2004). Glutamate release has been documented from midbrain neurons in culture, and cultured dopaminergic cells co-express both VGLUT2 and VMAT2 (Sulzer et al., 1998; Dal Bo et al., 2004). In vivo studies have yielded less consistent results. One group has reported that mRNA for VGLUT2 is likely to be expressed in dopaminergic neurons in the ventral tegmental area (Kawano et al., 2006). However, others report that VGLUT2 and VMAT2 mRNA localize to distinct cells (Yamaguchi et al., 2007). Furthermore, VMAT2 and VGLUT2 show minimal co-localization in dopaminergic nerve terminals (Kawano et al., 2006). Thus, it remains unclear whether dopamine and glutamate are co-released from midbrain dopaminergic neurons in vivo. If future studies do indeed validate the idea that dopamine and glutamate are co-released, it would have important ramifications for their reciprocal regulation in the striatum, as well as disease processes such as addiction and schizophrenia that are thought to involve interactions between dopaminergic and glutamatergic circuits (Zhang and Sulzer, 2003; Kalivas, 2004; Carlsson, 2006). In contrast to mammalians systems, in which two isoforms of VMAT, and three isoforms of VGLUT are expressed, Drosophila and C. elegans each express a single ortholog of VGLUT and VMAT (the genes dVGLUT (Daniels et al., 2004) and dVMAT (Greer et al., 2005) in Drosophila, and eat-4 (Avery, 1993; Bellocchio et al., 2000) and cat-1 (Duerr et al., 1999)) in C. elegans. In C. elegans, four motoneurons (VC4, VC5, HSNL, HSNR) express both the vesicular acetylcholine transporter and the vesicular monoamine transporter (Duerr et al., 2001). Indeed, this was the first reported example of neurons expressing two different vesicular neurotransmitter transporters in vivo (Duerr et al., 2001). Interestingly,

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immunohistochemical analysis has shown that the two transporters do not consistently co-localize at a subcellular level (Duerr et al., 2001). This observation suggests that different transporters might sort to different populations of SVs within the same neuron, as proposed for mammalian VGLUT1 and 2 (Fremeau Jr et al., 2004a). In Drosophila, DVMAT and DVGLUT are co-expressed in nerve terminals that release octopamine and presumably glutamate at Type II neuromuscular junctions (Greer et al., 2005). The mechanisms by which co-release of two transmitters may affect synaptic transmission are not clear. This is especially true for examples in which the co-released transmitters produce opposing post-synaptic effects, such as excitation and inhibition in the case of GABA and glutamate. Multiple hypothetical mechanisms may account for this interesting phenomenon, but to date, there is little or no experimental data. Finally it should be noted that some neurons in the adult express two vesicular transporters of the same type. In rat and mouse, only VMAT1 is expressed in the neurosecretory cells of the adrenal medulla for release of norepinephrine and epinephrine (Peter et al., 1995; Erickson et al., 1996). However, in primates, both VMAT1 and 2 are expressed in the same cells in the adrenal medulla (Weihe and Eiden, 2000). Both VMAT1 and 2 may also be expressed in small intensely fluorescent (SIF) cells of the sympathetic ganglia (Weihe and Eiden, 2000). As noted above, both VMAT1 and 2 are co-expressed in the adrenal medulla and SIF cells of the sympathetic and parasympathetic ganglia in some species (Schutz et al., 1998; Weihe and Eiden, 2000). The functional importance of VMAT co-expression is unclear, but could allow complimentary substrate affinities.

8

Biosynthesis and Trafficking

8.1 Biosynthesis As is the case for other integral membrane proteins, vesicular transporters are synthesized in the endoplasmic reticulum (ER), and post-translationally modified in the ER and Golgi. Based on their sensitivity to glycosidase and the results of mutagenesis studies, VMATs and VAChT undergo N-linked glycosylation (Tan et al., 1998; Yelin et al., 1998; Yao et al., 2004; Yao and Hersh, 2007). Mutation of three predicted glycosylation sites in VMAT1 decreases transport activity but does not affect substrate affinity (Yelin et al., 1998). Decreased glycosylation does not disrupt subcellular localization of VMAT1 in non-neuronal cells (Yelin et al., 1998) but may help mediate the localization of VMATs to LDCVs (Yao and Hersh, 2007) (see below). In addition to glycosylation, other potential modifications during protein biosynthesis include phosphorylation, covalent attachment of lipids, and formation of multimers. It is likely that phosphorylation of VMAT2 occurs prior to exit from the Golgi, since a partially glycosylated species likely to be an immature form of VMAT2 is phosphorylated by CKII (Krantz et al., 1997). There is very little additional biochemical information on vesicular transporter biosynthesis, and other biosynthetic events remain incompletely characterized, or are a matter of speculation. In particular, it remains possible that some vesicular transporters form multimers (Gracz et al., 1988), but there is little data supporting this possibility. In addition, based on their size in SDS-PAGE gels VGLUTs, but not VGAT, are likely to be glycosylated.

8.2 Membrane Trafficking The trafficking and subcellular localization of vesicular transporters depends primarily on signals encoded in their cytoplasmic domains. Most signals identified thus far are related to known, cytoplasmic trafficking motifs, which presumably bind to the machinery responsible for trafficking many other membrane proteins (for a general review of trafficking motifs, see (Bonifacino and Traub, 2003)). The trafficking machinery includes vesicle coat proteins, such as clathrin that help sequester membrane proteins into budding vesicles (Ungewickell and Hinrichsen, 2007). In addition, adaptor proteins such as the clathrin adaptor protein complexes (APs) function as intermediates to link the vesicular proteins to the coats (Boehm and Bonifacino, 2001; Dell’Angelica, 2001). A large number of accessory proteins associate with the adaptors and coats. Some, such as endophilin, may help remodel lipid membranes (Gallop et al., 2006). Together,

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these proteins help vesicular transporters exit the Golgi in the secretory pathway and allow their uptake from the plasma membrane in the endocytic pathway. Both secretory and endocytic pathways are required for vesicular transporter trafficking to secretory vesicles: the biogenesis of synaptic vesicles is thought to involve an endocytic step at the plasma membrane of the nerve terminal, and SV recycling also requires endocytosis (Hannah et al., 1999; Prado and Prado, 2002; Su¨dhof, 2004; Jahn and Rizzoli, 2007). However, these trafficking steps remain incompletely understood, and contribution of particular motifs for sorting vesicular transporters to SVs is not fully defined. With these caveats in mind, we review the progress in the field of vesicular transporter trafficking (see also our 2008 review in Traffic). For VMAT2, a number of studies indicate that the cytoplasmic Cterminal domain is responsible for endocytosis from the plasma membrane and sorting to both SVs and LDCVs (Erickson and Varoqui, 2000). A dileucine motif is required for efficient endocytosis of VMAT2 in PC12 cells as well as in hippocampal neurons (Tan et al., 1998; Li et al., 2005) and is likely to help sort VMAT to SLMVs in PC12 cells (Colgan et al., 2007; Yao and Hersh, 2007) (see > Figure 7-3a). However,

. Figure 7-3 Membrane trafficking signals. Glycosylation (‘‘Y’’) facilitates sorting of VMAT2 to large dense core vesicles (LVs). All other known sorting signals for vesicular transporters are encoded in their C-terminal, cytoplasmic domains. For VMAT2, a dileucine motif (IL) is required for endocytosis and sorting to synaptic like microvesicles (SLMVs) in neuroendocrine cells. The VMAT2 endocytosis signal is part of a larger motif, which includes upstream glutamate residues (EE). Upstream glutamates are required for localizing VMAT2 to LVs. A phosphorylated ‘‘acidic cluster’’ at the extreme C-terminus of VMAT2 (DDEE[P]SE[P]SD, shown as [P]SD in panel (a)) also helps sort VMAT2 to LVs. (b) VAChT contains a dileucine motif (LL) required for endocytosis and sorting to SLMVs. However, a tyrosinebased motif (Y) may direct these trafficking events under some circumstances. A serine upstream of the dileucine motif in VAChT (SE) undergoes phosphorylation by PKC. Phosphorylation of the serine ([P]SE) may drive a portion of VAChT onto LVs. (c) Endocytosis and recycling of VGLUT1 to SVs requires a dileucine-like motif (FV). Under some circumstances, a polyproline motif (PP) also contributes to VGLUT1 recycling to SVs

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since multiple forms of endocytosis may occur in neurons (Galli and Haucke, 2004; Wienisch and Klingauf, 2006), and proteins may sort to neuronal SVs via alternate mechanisms (Thoidis et al., 1998; Blagoveshchenskaya and Cutler, 2000), it is possible that other motifs could participate in the sorting of VMAT2 to SVs in vivo. Additional motifs are required for localization of VMAT2 to LDCVs. These include acidic glutamate (E) residues upstream of the dileucine motif (EEXXXIL) (Dietrich et al., 1994; Pond et al., 1995; Krantz et al., 2000). In addition, an acidic cluster or patch at the end of the C-terminal (DDEESESD) domain helps traffic VMAT2 to LDCVs (Waites et al., 2001). This motif was originally identified in the protease furin (Wan et al., 1998), and for both VMAT2 and furin, the acid patch may determine whether protein is retained in LDCVs as they mature (Scott et al., 2006). N-linked glycosylation may provide a third signal for localizing VMAT2 to LDCVs in PC12 cells. The glycosylated loop is not itself sufficient to target VMAT2 to LDCVs. However, the C-terminal domain of VMAT2 is only competent for sorting to LDCVS if the lumenal loop is also present. Moreover, pharmacologic blockade of N-linked glycosylation prevents VMAT2 from sorting to LDCVs (Yao and Hersh, 2007). Potentially related phenomena include the observation that glycosylation is decreased if VMAT2 is mislocalized to SLMVs in PC12 cells (Yao et al., 2004). In addition, a decrease in VMAT2 glycosylation with increasing age in rats may correlate with a decrease in its localization to synaptic vesicles (Cruz-Muros et al., 2008). For VAChT, endocytosis may depend on a dileucine motif similar to a site in VMAT2 (Tan et al., 1998; Santos et al., 2001; Barbosa et al., 2002) or a distinct, downstream tyrosine-based motif (Kim and Hersh, 2004). Consistent with the predicted role for endocytosis in SV biogenesis, mutation of the dileucine motif blocks the localization of VAChT to SLMVs in PC12 cells (Colgan et al., 2007). VAChT also contains an upstream acidic residue and a PKC phosphorylation site that can mimic acidic residues in the extended dileucine motif of VMAT2 ([Phospho-S]EXXXLL) (Cho et al., 2000; Krantz et al., 2000). Furthermore, substitution of an acidic amino acid at this site drives VAChT to LDCVs in PC12 cells, mimicking the trafficking pattern of VMAT2 (Krantz et al., 2000). However, phosphorylation by PKC is not constitutive and VAChT lacks the additional acidic patch found in VMAT (Krantz et al., 2000). It is likely that these differences contribute to the divergent localization of VMAT2 and VAChT in PC12 cells, in which VAChT localizes primarily to SLMVs, and VMATs primarily to LDCVs (Liu et al., 1994; Liu and Edwards, 1997). Similarly, in cholinergic neurons in vivo, VAChT localizes primarily to SVs (however, see (Lundberg et al., 1981; Agoston and Whittaker, 1989; Krantz et al., 2000)). As for VMAT2 and VAChT, a dileucine-like motif in VGLUT1 appears to play an important role in endocytosis, and presumably its localization to SVs (Voglmaier et al., 2006). However, at least two modes of endocytosis have been proposed for VGLUT1 (Voglmaier et al., 2006). In one, the dileucine-like motif mediates rapid endocytosis, most likely through the clathrin-dependent adaptor complex AP2. Under some circumstances, a polyproline motif present in VGLUT1 (but absent from other VGLUT isoforms) works in concert with the dileucine-like motif. A slower mode of VGLUT endocytosis has been proposed to be mediated by the adaptor protein complex AP3 (Voglmaier et al., 2006). In a separate set of experiments, another group has shown that localization of VGLUT to SLMVs is likely to depend on the adapter protein AP3 in PC12 cells (Salazar et al., 2005). It should be noted that the relative contribution of AP3-dependent trafficking to SV biosynthesis in neurons is controversial (Mullins et al., 2000). Nonetheless, these results indicate that the localization of VGLUTs to SVs may involve at least two distinct trafficking pathways. In future experiments, it will be critical to determine how each pathway contributes to regulation of glutamate release in vivo. In a few cases, vesicular transporters have been shown to directly bind adaptor proteins and other elements of the trafficking machinery. VGLUT1 binds endophilin (Voglmaier et al., 2006) and VMAT2 binds the adaptor PACS-1, possibly regulating exit from immature LDCVs (Waites et al., 2001). VAChT has been suggested to bind AP1 and AP2 (Barbosa et al., 2002; Kim and Hersh, 2004). In addition to these wellknown adaptor proteins, it is possible that vesicular transporters require other proteins to interact with the sorting machinery. In C. elegans, a screen for genes that suppressed the phenotype of a VAChT mutant (unc17(e245)) identified a mutant allele of synaptobrevin (snb-1) (Sandoval et al., 2006). Results of additional genetic experiments support a direct interaction between the mutant alleles of VAChT and synaptobrevin

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(Sandoval et al., 2006). In addition to rescuing the behavioral phenotype, the snb-1 allele also rescued a decrease in VAChT expression caused by the unc-17(e245) mutation (Sandoval et al., 2006). Although the function of the VAChT/synaptobrevin interaction is not known, it is tempting to speculate that it could facilitate localization of VAChT to SVs. The trafficking of VGAT has only recently been investigated and sorting signals have not yet been defined. However, based on a decrease in GABAergic neurotransmission in knockout mice lacking the mu subunit of the clathrin adaptor protein complex AP3, it has been proposed that AP3 may help sort VGAT to SVs (Nakatsu et al., 2004). The mouse knockout of the mu3B subunit of the neuron-specific AP3B complex shows decreased GABAergic neurotransmission. In addition, the density of synaptic vesicles in inhibitory terminals was lower in 3B knockout mice than in wild-type mice, suggesting that AP-3B is involved in the biogenesis of GABA containing synaptic vesicles. A small change was also seen in the localization of VGAT to endosomes, although immunolabeling did not detect mislocalization of VGAT. Additional experiments will be needed to definitively determine whether AP3 is directly involved in VGAT trafficking. Genetic studies in C. elegans have shown that sorting of VGAT to SVs may require the novel protein unc-46 (Schuske et al., 2007). Mechanisms underlying their interaction are not yet known. Nonetheless, this finding is important since it suggests that specialized proteins acting in concert with the standard trafficking machinery may be required for sorting vesicular transporters to secretory vesicles. We anticipate that future genetic screens in both C. elegans and Drosophila may help identify additional elements specialized for trafficking vesicular transporters. Unlike VGLUT 1 and 2, VGLUT3 is expressed in dendrites as well as axon terminals (Fremeau et al., 2002, 2004b). The dendritic localization of VGLUT3 corresponds to vesicular structures that appear to mediate regulated release (Harkany et al., 2004). These vesicles presumably differ from synaptic vesicles since other markers for SVs are not found in dendrites, and dendrites do not contain active zones for SV release. Regardless of the precise vesicle type, the possibility of regulated glutamate release from dendritic sites has very important ramifications for signaling. Indeed, VGLUT3-dependent glutamate release has been implicated in retrograde signaling from post-synaptic sites in cortical pyramidal cells (Zilberter et al., 1999; Harkany et al., 2004). Interestingly, GABA release from dendrites also has been reported to mediate retrograde signaling in the cortex (Zilberter et al., 1999). It is therefore possible that VGAT might localize to analogous structures in GABAergic neurons, and release GABA in a manner analogous to dendritic glutamate release by VGLUT3. In some cases, transporter expression may influence SV trafficking and/or release. Neither VMAT2 (Croft et al., 2005) nor VAChT knockout mice (Prado et al., 2006) shows any apparent defects in SV recycling. Similarly, depletion of acetylcholine via inhibition of VAChT activity does not affect trafficking of cholinergic vesicles (Parsons et al., 1999). In addition, application of bafilomysin to deplete proton gradient does not alter SV recycling in glutamatergic neurons, despite a dramatic reduction in vesicular glutamate (Zhou et al., 2000). In contrast, loss of VGLUT1 may result in aberrant membranous structures possibly reflecting alterations in endocytosis (Fremeau et al., 2004a). It is also possible that loss of VGLUT1 alters the subcellular distribution of SVs in glutamatergic terminals. Most nerve terminals contain at least two functionally distinct pools of SVs, including those that cluster close to the plasma membrane and are ‘‘readily released.’’ SVs in a distinct ‘‘reserve pool’’ undergo fusion only after the readily releasable pool has been depleted. Electron micrographs of glutamatergic nerve terminals in VGLUT1 knockout mice show wild type numbers of SVs immediately adjacent to the plasma membrane, presumably consistent with an intact readily releasable pool. However, the number of vesicles more distant from the membrane is depleted, suggesting a reduction in the reserve pool (Fremeau et al., 2004a). VGLUT1 heterozygotes show a similar pattern, consistent with a reduction of the reserve pool (Tordera et al., 2007). It is possible that reduction in the amount of glutamate in an individual SV could alter its localization, or a decrease in glutamate release could cause a reduction of the reserve pool via a more indirect, feedback mechanism. However, as noted above, recycling of SVs is not altered by depletion of glutamate (Zhou et al., 2000). Additional experiments will be needed to determine how VGLUTs may influence SV trafficking.

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8.3 How Many Transporters are on Each Vesicle? In our discussion of transporter biosynthesis and trafficking, we have glossed over the question of how many vesicular transporters reside on each synaptic vesicle. Surprisingly, this question has not yet been resolved. For VAChT, the Hill coefficients for vesamicol binding suggest that it might be an oligomer (Gracz et al., 1988). More recently, quantitative mass spectroscopic analysis of SVs suggested that 10–14 VGLUTs are associated with each vesicle (Takamori et al., 2006). Aside from these results, most data addressing this question are derived from electrophysiological studies. Knockout mice and transporter mutants have provided a critical experimental platform for these experiments. It may be argued that if more than one transporter resides on each wild type vesicle, reducing transporter expression should result in a mixed population of vesicles, with varying numbers of transporters and thereby, varying transmitter content. This can be assayed using frequency and amplitude of miniature excitatory potentials (minis), which represent post-synaptic response to a single vesicle (> Figure 7-4). A decrease in mini frequency

. Figure 7-4 How many transporters are on each vesicle? The question of how many vesicles reside on each vesicle many be assayed electrophysiologically. The cartoon shows how quantal size, the electrophysiologic response to a single SV, may be used to determine whether a vesicle uses 1 versus  2 transporters to load baseline levels of neurotransmitter. Quantal size is reflected by the post-synaptic response to a single vesicle, e.g., a miniature or ‘‘mini’’ excitatory potential (mEPSP). If most vesicles use  2 transporters it would be expected that a decrease in expression would cause a decrease in the amount of transmitter that is loaded into each vesicle. In contrast, if most vesicles use only one transporter, then a decrease in expression is likely to lead to some vesicles lacking a transporter, and therefore devoid of transmitter. This will be reflected in a decrease in the frequency of minis that are observed. However, with low but detectable levels of expression, a few vesicles may have a single transporter. If this is the same number of transporters on most vesicles under baseline conditions the few minis that are observed will not show a detectable decrease in size

suggests an increase in the number of empty vesicles. In contrast, a decrease in mini amplitude may be interpreted as evidence for a decrease in the amount of neurotransmitter stored in an individual SV, excluding any post-synaptic changes. If more than one transporter localizes to each SV, a decrease in transporter expression would be expected to result in a decrease in the number of transporters on each

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vesicle, and thus a reduction in mini size (> Figure 7-4, top panel). Thus, in the simplest case in which there are two transporters on each vesicle, there would be two classes of detectable mini amplitudes when transporter expression is reduced. The larger minis would represent SVs with two transporters and the smaller class would represent SVs with only one transporter.2 Conversely, if only one transporter resides on each vesicle, then reducing transporter expression should result in some vesicles without a transporter (> Figure 7-4, bottom panel). Exocytosis of empty vesicles should not result in a postsynaptic response, which can be seen as a decrease in mini frequency. However, the remaining minis should show an average amplitude identical to wild type, since both have a single vesicular transporter. Within this conceptual framework, results from Drosophila, zebrafish and mice appear to yield conflicting results. In Drosophila, dVGLUT mutants expressing a small but detectable level of DGLUT, most vesicles appeared to be empty, judging by a decrease in the frequency in the number of minis (Daniels et al., 2006). Surprisingly, the remaining minis were equivalent to wild type in amplitude suggesting that the amount of glutamate in each vesicle was equivalent (Daniels et al., 2006). In contrast, results from a Zebrafish VGLUT2 mutant suggest that more than one transporter may reside on single vesicle under some circumstances, since the mutants showed a reduction in the mean mEPSC amplitude (Smear et al., 2007). VGLUT1 knockout mice also have been used to address this question, but results vary across laboratories and preparations. One group used primary cultures of glutamatergic neurons that co-expressed VGLUT1 and 2 (Wojcik et al., 2004). Primary cultures from VGLUT1 knockout homozygotes showed a decrease in mini amplitude, indicating decrease in quantal size (Wojcik et al., 2004). Similar results were seen using cultured neurons from VGLUT1 heterozygotes (Tordera et al., 2007). The most likely explanation for these results is that both VGLUT1 and 2 can reside on a single vesicle in wild type cells. In the absence of VGLUT1, only VGLUT2 would be available to package glutamate, and the vesicle would be partially filled. In contrast to these results, another group did not detect a decrease in mini size in organotypic slice-based experiments using tissue from VGLUT1 knockout mice (Fremeau et al., 2004b). These results were interpreted as indicating that that VGLUT1 and 2 reside on distinct and mutually exclusive populations of SVs (Fremeau et al., 2004c). The most significant technical difference between these two sets of experiments is that the group showing a decrease in quantal size used primary neuronal cultures, whereas the group that did not detect any differences used a slice preparation. It is possible that neurons in the slice experiments had a more mature phenotype, and expressed either VGLUT1 or VGLUT2 but not both, consistent with expression pattern in adult animals (Boulland et al., 2004). Conversely, at least in some hippocampal neurons early in development, VGLUT1 and 2 can localize to the same vesicle (Herzog et al., 2006). In sum, available data does not yet definitively establish whether one or more than one transporter resides on each SV. However, as we discuss below, calculations to determine the rate of vesicle filling make it difficult to understand how a single transporter can suffice to fill a vesicle within the time thought to be required for recycling. Thus, it may be more reasonable to consider a ‘‘single’’ transporter to be composed of an indivisible cluster of several transport proteins. While attractive from a physiological standpoint, the only biochemical data of which we are aware indicating that vesicular transporters form oligomeric structures is the positive Hill coefficient of vesamicol binding to VAChT (Gracz et al., 1988).

9

Regulation of Expression and Activity

9.1 Transcription Since changes in the number of transporters per vesicle can alter transmitter release, transcriptional regulation of transporter expression may have a significant impact on signaling. In the periphery,

2

We note that several assumptions must be made in this admittedly over-simplified scenario. For example, for a net increase in neurotransmitter transport to occur, it must be assumed that an increase in inward flux is not negated by outward ‘‘leakage’’ of transmitter through the same transporter.

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transcriptional regulation of VMAT2 expression has been studied intensively in an effort to understand the regulated release of histamine in the gut. In the gastric mucosa of humans and rodents, histamine is released from secretory vesicles in enterochromaffin-like (ECL) cells, and histamine release in turn regulates gastric acid secretion in the stomach. Coordinate regulation of VMAT2 and the biosynthetic enzyme histidine decarboxylase (HDC) may represent an important mechanism of controlling acid secretion (Dockray et al., 2001). The regulation of VMAT2 transcription has been demonstrated in cultured cell lines including the gastric epithelial cell line AGS-G R (Watson et al., 2000). These cells express the gastrin-CCK beta receptor, and stimulation with the hormone gastrin increases transcription of VMAT2 mRNA (Watson et al., 2000). Gastrin also increases expression of HDC in enterochromaffin cells (Dimaline and Sandvik, 1991; Zhang et al., 1996; Hocker et al., 1997). An increase in mRNA for both HDC and VMAT2 has been observed in animal models of hypergastrinemia, suggesting that transcriptional activity of these genes may be regulated by gastrin in vivo (Dimaline and Sandvik, 1991, 1996). The mechanism by which VMAT2 expression is regulated by gastrin may involve a novel function for the proteasome (Catlow et al., 2007). A proteasomal subunit was shown to bind a ten base-pair segment in the VMAT2 promoter and thereby enhance gastrin-induced secretion (Catlow et al., 2007). Importantly, this activity is independent of the degradative functions of the proteasome elsewhere in the cell (Catlow et al., 2007). It will be interesting to see if this represents a general mechanism for regulating VMAT and whether it occurs in the central nervous system as well as the gut. A number of other studies have investigated the transcriptional regulation of VMAT2 in neurons and neuroendocrine cells. Early studies using primary cultures of chromaffin cells and sympathetic ganglia suggested that increased stimulation and calcium influx might up-regulate VMAT transcription (Desnos et al., 1990, 1992; Krejci et al., 1993). Conversely, minimal changes in expression were seen in vivo in response to VMAT inhibition using reserpine (Vilpoux et al., 2000). Since most psychiatric illnesses have been linked in one way or another with monoamines, there has been a great deal of interest in establishing a link between VMAT expression and other drugs that are used in clinical psychiatry. In rats, chronic administration of either the antipsychotic clozapine (Rehavi et al., 2002) or the mood stabilizer lithium (Cordeiro et al., 2002) increases VMAT2 expression, presumably via increased transcription. VAChT is encoded in the first intron of the biosynthetic enzyme for ACh, ChAT, and is in the same transcriptional orientation. This peculiar relationship is evolutionarily conserved from C. elegans to humans suggesting an important functional role, and the possibility of co-regulated gene transcription (Alfonso et al., 1994; Bejanin et al., 1994; Erickson et al., 1994; Roghani et al., 1994; Kitamoto et al., 1998). Indeed, studies in cultured neurons indicate that expression of ChAT and VAChT are coordinately upregulated by retinoic acid and leukemia inhibitory factor/ciliary neurotrophic factor (LIF/CNTF) (Berrard et al., 1995; Berse and Blusztajn, 1995; Misawa et al., 1995). An upstream neuron-restrictive silencer element (NRSE) may function to coordinately regulate expression of both genes (Shimojo et al., 1999; De Gois et al., 2000). However, under some circumstances VAChT and ChAT may be differentially expressed. In mammals, multiple regulatory elements are involved in VAChT and ChAT transcription including promoter elements for VAChT contained within the ChAT intron (Berse and Blusztajn, 1995; Cervini et al., 1995; Schutz et al., 2003). Some stimuli such as cAMP differentially affect ChAT versus VAChT expression in vitro (Berse and Blusztajn, 1995; Misawa et al., 1995) and in vivo - the ratio of VAChT versus ChAT transcript is higher in peripheral versus central cholinergic neurons (Schutz et al., 2001). In addition, ChAT but not VAChT is expressed in the placenta (Oda et al., 2004). Some of the genomic regulatory regions responsible for the differential expression of ChAT and VAChT have been mapped using transgenes containing varying amounts of the putative regulatory sequences; at least one segment of genomic DNA was shown to control expression of ChAT but not VAChT (Schutz et al., 2003). Dissociation of ChAT and VACHT transcription is highlighted by the analysis of VAChT knockdown mice (Prado et al., 2006) in which ChAT expression is unchanged (Prado et al., 2006). The mechanisms regulating VGLUT expression have yet to be determined. However, upregulation of VGLUT has been observed in rat cortex and hippocampus in response to antidepressant and electroconvulsive shock treatment (Moutsimilli et al., 2005; Tordera et al., 2005). This data suggests the possibility that VGLUTs undergo transcriptional regulation in vivo (see also > Section 9.6).

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9.2 Alternative mRNA Splicing Alternative splicing has been proposed to regulate the function of several plasma membrane transporters including GLYT-1 (Borowsky and Hoffman, 1998; Hanley et al., 2000; Poyatos et al., 2000), and could potentially regulate the function of vesicular neurotransmitter transporters as well. Both VAChT and ChAT undergo alternative splicing of 50 non-coding regions, and the splicing of ChAT and VAChT might be coordinated, although the functional significance of these splicing events are not yet clear (Bejanin et al., 1994; Cervini et al., 1995). A VGAT splice variant has been reported in pancreatic tissue, and shows a predicted amino-terminal truncation (Suckow et al., 2006). Another proposed VGAT variant retains intron 2 as an exon (Ebihara et al., 2003) but may represent a cloning artifact (Oh et al., 2005) and the possibility of alternative splicing yielding a functional divergent protein remains an open question in mammalian systems. In Drosophila, DVMAT shows variable use of a 30 splice site in the last exon of the gene, which leads to two divergent carboxy-terminal domains (Greer et al., 2005). As a result of this difference, only one version undergoes efficient endocytosis in vitro (Greer et al., 2005). Since endocytosis is thought to be required for localizing vesicular transporters to synaptic vesicles, this data suggests that alternative splicing could regulate trafficking of DVMAT to secretory vesicles.

9.3 Trafficking and Transporter Phosphorylation VMATs, and possibly VAChT, localize to both SVs and LDCVs. SVs cluster at active zones in the nerve terminal and mediate rapid signaling. In contrast, large dense core vesicles (LDCVs) occur diffusely throughout the cell body and dendrites as well as in axon terminals. LDCVs respond to different stimuli than synaptic vesicles, and also co-release peptide neurotransmitters (Kelly, 1993). Since VMAT and VAChT localize to both vesicle types, regulated changes in transporter trafficking has the potential to dramatically change the mode of neurotransmitter release. In addition, since the absolute number of transporters that reside on each secretory vesicle can directly regulate transmitter storage, transporter trafficking has the potential to increase the amount of transmitter that is released from both types of vesicle. Trafficking of both VAChTand VMAT2 is directly regulated by phosphorylation of cytoplasmic trafficking domains. For VAChT, a serine five residues upstream of the dileucine motif is phosphorylated by PKC (Cho et al., 2000; Krantz et al., 2000). A distinct C-terminal cluster of acidic residues and serines in VMAT2 is phosphorylated by CKII (Krantz et al., 1997). For both transporters, substitution of acidic residues at the phosphorylation sites has been used to assess the potential effects of phosphorylation on transporter trafficking (Krantz et al., 2000; Waites et al., 2001). Phosphorylation of VAChT by PKC converts the serine to an acidic phosphoserine similar to the acidic glutamate residue at the analogous position in VMAT2. Substitution of a glutamate for the serine in VAChT increased sorting of VAChT approximately threefold in PC12 cells (Krantz et al., 2000). It is likely that phosphorylation at this site affects trafficking at the TGN, and the sorting of VAChT into the constitutive versus regulated secretory pathway. However, it remains possible that phosphorylation of VAChT regulates other trafficking events. Although mutation of sites upstream of the dileucine motif does not appear to effect transporter endocytosis in cultured cell lines (Tan et al., 1998) phosphorylation of VAChT in neurons could potentially regulate trafficking at the synapse. In hippocampal slices, activation of PKC blocks the ability of the VAChT inhibitor vesamicol to inhibit acetylcholine release, and this effect correlates with an increase in VAChT phosphorylation (Barbosa et al., 1997). For VMAT2, an additional acid patch motif at the extreme C-terminus also contributes to its localization to LDCVs (Waites et al., 2001). The substitution of acidic residues for serines imbedded in the acid patch motif the increases localization of VMAT2 to LDCVs (Waites et al., 2001). Unlike most kinases, CKII is constitutively active and VMAT2 is therefore phosphoryated by CKII under baseline conditions (Krantz et al., 1997; Pinna, 2002). Moreover, since an immature VMAT2 species is phosphorylated, CKII is most likely to act before exit of VMAT2 from the Golgi, consistent with a role for phosphorylation in regulating VMAT2 trafficking at the TGN. Similar to furin - which also undergoes phosphorylation by CKII (Wan et al., 1998)- this is likely to determine whether VMAT2 is retained in LDCVs as they mature at the TGN. This raises the possibility that dephosphorylation of VMAT2 and removal from maturing LDCVs could

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increase VMAT2 sorting to other compartments, and perhaps increase its localization to SVs at the synapse. To test this possibility, it will be useful to study VMAT trafficking in vivo. VGAT also undergoes phosphorylation, and interestingly, phosphorylation of VGAT has been observed in neurons, but not other non-neuronal cells (Bedet et al., 2000). This difference suggests the possibility of a specific neuronal function for VGAT phosphorylation, but this remains untested. It is also unknown whether direct phosphorylation regulates the function of VGLUT, or indeed if it is phosphorylated at all. Phosphorylation may also indirectly regulate transporter function. Additional regulatory events associated with phosphorylation are suggested by a series of physiological studies on cholinergic signaling at the NMJ performed in the Van Der Kloot lab. These show that during periods of sustained SV release, PKC decreases (Van der Kloot, 1991) and PKA increases quantal size (Van der Kloot and Branisteanu, 1992). Further studies will be needed to elucidate the mechanisms underlying these phenomena, and their relationship to VAChT phosphorylation and trafficking. In addition, although PKA does not directly phosphorylate VMATs, it is required for the localization of VMAT1 and 2 to LDCVs in PC12 cells (Yao et al., 2004).

9.4 Heterotrimeric G Proteins Transporters also may be directly regulated via protein-protein interactions or post-translational modifications that increase or decrease transport activity. A series of papers from the Ahnert-Hilger lab over the past 10 years suggest that VGLUT and VMAT activity are regulated by heterotrimeric G proteins (Brunk et al., 2006b). This unexpected finding was first reported in 1998 using PC12 cells, which endogenously express VMAT1 (Ahnert-Hilger et al., 1998) and subsequent work has begun to elucidate the underlying mechanisms (Brunk et al., 2006b) (> Figure 7-5a). Vesicular transporters interact with previously identified . Figure 7-5 Vesicular transporters may undergo multiple forms of regulation. These include: (a) Regulation of VMAT2 activity by vesicular G-proteins; (b) Regulation by other directly associated proteins. VGAT binds to glutamic acid decarboxylase (GAD), which may help couple synthesis and transport of GABA. GAD and perhaps VGAT are part of a larger protein complex that includes cysteine string protein (CSP) and the heat shock protein HSC70; (c) Post-translational regulation of VGLUT1 expression by the clock protein Per (see text for details)

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G proteins, but the signaling mechanisms used by transporters versus G protein coupled receptors differ; most importantly, interactions with both VMATs and VGLUT are likely to occur on secretory vesicles rather than cell surface (Pahner et al., 2002, 2003). There, VMAT1 and 2 can interact with either Galpha(o2) or Galpha(q) depending on cell type (Holtje et al., 2000, 2003). In each case, the downstream effect is inhibition of VMAT activity (Ahnert-Hilger et al., 1998; Holtje et al., 2000, 2003). Current models suggest that a lumenal domain in the transporter itself may activate the G protein and that binding of substrate to VMAT is required for G protein inhibition of VMAT activity (Brunk et al., 2006a). This in turn suggests that the function of this regulatory process may be to limit transport, as the vesicle lumen fills with neurotransmitter (> Figure 7-5a). The manner in which G proteins inhibit VMAT activity and the requirement for additional proteins remains unclear (Brunk et al., 2006a). Similar to VMATs, all three mammalian VGLUT isoforms have been proposed to undergo regulation by the heterotrimeric G proteins Galpha(o2) (Winter et al., 2005). In the absence of Galpha(o2), the usual biphasic dependence on Cl concentrations is essentially abolished (Winter et al., 2005). It has been suggested that regulation by Galpha(o2) might serve to control glutamate release in response to increased levels of cellular activity (Winter et al., 2005).

9.5 Protein Binding Partners (see also > Section 8.2) Additional proteins proposed to interact with vesicular transporters include enzymes responsible for neurotransmitter biosynthesis. The idea that biosynthetic enzymes interact with the transporters is attractive in that it would help explain the long-standing observation that newly synthesized transmitter is preferentially transported into SVs (Besson et al., 1969; Collier, 1969; Jin et al., 2003). Coupled synthesis and transport has been suggested for dopamine (Chen et al., 2003), acetylcholine (Sha et al., 2004) and GABA (Jin et al., 2003). However, the most convincing evidence to date suggests that biosynthetic enzyme for GABA, L-glutamic acid decarboxylase (GAD) forms a complex with VGAT and at least three other proteins including Cysteine String Protein (CSP), calcium calmodulin kinase II (CamKII) and heat shock cognate 70 (HSC70) (Jin et al., 2003) (> Figure 7-5b). If borne out by additional experiments, these relationships suggest that neurotransmitter synthesis and packaging, similar to other cellular processes, may occur on a complex or scaffold that could facilitate the packaging process itself, as well as coordinate regulation of its components. It is tempting to consider the possibility that transmitter uptake at the plasma membrane could also be physically linked to vesicular transport. An additional protein proposed to bind to VGLUT and VGAT has been designated inhibitory protein factor (IPF) (Ozkan et al., 1997; Tamura et al., 2001). IPF is a cytosolic fragment of the cytoskeletal protein fodrin, and interactions with VGAT and VGLUT may decrease the exocytotic release of both GABA and glutamate. (Ozkan et al., 1997; Tamura et al., 2001). More recently, rat VAChT was found to directly interact with SEC14L1, which is thought to function as a phospholipid transfer protein (Ribeiro et al., 2007). SEC14L1 also binds to the high affinity choline transporter (CHT1), but not other SV proteins (Ribeiro et al., 2007). All of these interactions should occur on the cytosolic side of the membrane. In contrast, VAChT derived from the electric organ of the Torpedo fish immunoprecipitates with proteoglycan, suggesting an interaction with an extracellular site (Bahr et al., 1992b). It is conceivable that interaction of VAChT with proteoglycan could regulate the function of the transporter, thus presenting an interesting scenario in which localization to the cell surface could be coupled to changes in transport activity.

9.6 Upstream Regulatory Pathways Upstream events that control the expression and function of vesicular neurotransmitter transporters in the CNS are arguably the least understood aspect of their regulation. However, a few limited clues are available regarding cellular events and circuits that may be involved in these processes.

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Regulatory changes in VMAT trafficking have been proposed to occur in response to psychostimulants (Fleckenstein et al., 2007). Agents that block amine uptake at the plasma membrane such as cocaine and buproprion (Wellbutrin), increase the localization of VMAT to synaptic vesicles (Brown et al., 2001; Rau et al., 2005). In contrast, drugs that promote efflux such as amphetamines decrease the localization of VMAT to SVs (Brown et al., 2002). It has been suggested that the mechanism underlying this regulatory event involves upstream activation of dopamine autoreceptors by extracellular transmitter (Sandoval et al., 2002; Truong et al., 2004). However, the manner in which dopamine receptor stimulation regulates VMAT trafficking is not known. Furthermore, since cocaine and amphetamine both increase extracellular DA, it is unclear how they could have opposing effects on the localization of VMAT2 (Hanson et al., 2004). VGLUT1 protein levels have been shown to follow a circadian pattern that does not depend on transcription (Yelamanchili et al., 2006). This effect is abrogated by loss of the gene Period 2, which encodes a component of the circadian clock (> Figure 7-5c). Together, these data suggest that the output of upstream pacemaker cells somehow regulates the activity of VGLUT1 (Yelamanchili et al., 2006). Additional upstream mechanism are no doubt involved in the regulation of these and other vesicular transporters, and identifying these mechanisms remain one of the current challenges for understanding their role at the synapse.

10

Functional Effects of Altered Expression and Activity

10.1 Regulation of Quantal Size The response to transmitter released by a single vesicle is designated as quantal size. This was long considered to be determined exclusively at postsynaptic sites, since the amount of transmitter in a single vesicle was thought to be invariant. More recently, the idea that neurotransmitter content in synaptic vesicles can vary has gained acceptance in part due to accumulated evidence that post-synaptic receptors are not saturated. These include studies using hippocampal cultures and slices (Bekkers et al., 1990; Liu and Tsien, 1995; Mainen et al., 1999; McAllister and Stevens, 2000), the mossy fiber synapse of cerebellar granule cells (Silver et al., 1996), the calyx of Held (Yamashita et al., 2003), and additional GABAergic synapses (Barberis et al., 2004; Frerking et al., 1995). Simultaneous with the exploration of post-synaptic saturation, – molecular the genetic analysis of vesicular transporters has shown that presynaptic changes in their expression can regulate neurotransmitter release and post-synaptic signaling. The effects of both increased and decreased expression have been investigated. The first study to explore the effects of transporter over-expression employed VAChT and cocultures of frog neurons and myocytes (Song et al., 1997). Cholinergic signaling in this system was increased by the over-expression of the VAChT (Song et al., 1997). More recently, over-expression of VGLUT1 has been shown to potentiate quantal size in hippocampal cultures (Wojcik et al., 2004; Wilson et al., 2005) and at the glutamatergic NMJ in Drosophila (Daniels et al., 2004). It has not yet been directly demonstrated that the over-expression of VMAT can increase post-synaptic signaling. However, monoamine release can be directly measured using amperometry (Wightman et al., 1991). In this technique, amines are oxidized on a carbon fiber electrode to produce a measurable current. Amperometry has been used to show that over-expression of VMAT2 increases monoamine release from cultured neuroendocrine cells and dopaminergic neurons (Pothos et al., 2000). In addition, over-expression of Drosophila VMAT in vivo causes an increase in amine-dependent behaviors, consistent with an increase in a vesicular release (Chang et al., 2006). In vitro studies using psychostimulants support the idea that reducing vesicular amines can decrease quantal size (Sulzer et al., 1995). In the case of VGLUTs, the most direct support for an increase in quantal size has been an observed increase in mini size (Daniels et al., 2004; Wojcik et al., 2004; Wilson et al., 2005). In contrast the effect of increased minisize on a summed evoked response (representing many minis) has been less consistent across species; in mice, an increase in excitatory potentials was observed following over-expression of VGLUT1 (Wojcik et al., 2004; Wilson et al., 2005), but in flies overexpression of DVGLUT did not cause an increase in evoked potential (Daniels et al., 2004). This was suggested to be the result of homeostatic regulation of

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glutamatergic signaling at the fly NMJ. The question remains as to why similar compensatory effects were not seen in cultured neurons from the mouse (Wojcik et al., 2004). As discussed above (see > Section 8.3) loss of transporter function may decrease quantal size in VGLUT knockouts. VAChT knockdown mice also show a decrease in mini size in both homozygotes and heterozygotes (Prado et al., 2006) and VGAT homozygotes show a similar decrease in mini size (Wojcik et al., 2006). In sum, both increasing and decreasing transporter expression can affect post-synaptic signaling, but functional outcomes appear to vary. Further studies will be needed to determine whether these differences result from the relative maturity of the synapse, the use of intact synapses versus cultured neurons, or differences between mammalian and invertebrate systems. All of the studies described above involve artificial manipulation of transporter expression to increase or decrease quantal size. Is it possible that changes in vesicular transport could alter quantal size under physiological conditions? Under conditions of increased transporter expression this is certainly possible and, at least in the periphery, VMAT2 expression can be hormonally upregulated by gastrin (see above). Trafficking may also function to increase quantal size by adding more transporters to each vesicle. At the frog NMJ, presynaptic changes in acetylcholine release correlate with increased synaptic activity (Naves and Van der Kloot, 2001; Wang et al., 2005). Under these conditions, it is conceivable that transporters might be added to SVs in the endocytic pathway. In chromaffin cells, quantal size can rise with increased stimulation (Elhamdani et al., 2001). These vesicles do not recycle at the plasma membrane but it is possible that newly synthesized vesicles could contain additional transporters via trafficking events at the TGN (see above (Krantz et al., 2000; Waites et al., 2001)). In other cases the G protein mediated mechanisms described above may increase quantal size by increasing transport activity. Importantly, most of the studies described above have been performed in vitro, making it difficult to determine whether changes in quantal size could affect downstream processes and/or behavior. In contrast, a recent study in Drosophila investigated the relationship of quantal size at the NMJ to larval foraging behavior (Steinert et al., 2006). Under conditions in which food was limited, foraging larva that increased locomotion also showed an increase in glutamatergic transmission at the NMJ. The first component of this response was an increase in both mini size and synaptic vesicle diameter. Interestingly, larger vesicles that contained more glutamate were recruited from the reserve pool of vesicles, which are replenished via a endocytic mechanism distinct from that of the recycling pool (Kidokoro et al., 2004). Mutations in several vesicular and endocytic proteins have been shown to alter vesicle size (Zhang et al., 1998; Shimizu et al., 2003; Dickman et al., 2005; Poskanzer et al., 2006), and it is possible that these proteins also help determine the size of the reserve pool vesicles. It is possible that the reserve pool also has higher levels of DVGLUT. Importantly, if the reserve pool vesicles contain more DVGLUT than vesicles in the readily releasable pool, it would imply that endocytic trafficking of VGLUT to these divergent pools might be regulated. We anticipate that this system will be useful to further investigate potential role of vesicular transporters in regulating quantal size in vivo.

10.2 Cytoplasmic Clearance of Dopamine and Parkinson’s Disease Vesicular transport activities are clearly required for the storage and release of neurotransmitter from secretory vesicles. The notion that they may also regulate cytosolic levels of transmitter has gained attention more recently, particularly as a biologically relevant function for VMATs. Based on their similarity to bacterial transporters and their sequestration of MPP+, it has been suggested that VMATs might help detoxify cells (Schuldiner et al., 1995). In addition to exogenous toxins, endogenous monoamines and in particular dopamine may cause intracellular toxicity (Stokes et al., 1999). Although the precise mechanisms remain unclear, cellular damage due to dopamine may be the result of oxidative metabolites and/or more specific binding of dopamine to proteins directly linked to familial forms of Parkinson’s disease (Conway et al., 2001; LaVoie et al., 2005). The regulation of cytosolic dopamine by VMAT and potential ramifications for neuroprotection has been the subject of several studies, both in vitro and in vivo. In vitro overexpression of VMAT2 in PC12 cells

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decreases cytosolic DA, as shown indirectly by decreased neuromelanin formation, and more directly by amperometric measurements of cytoplasmic amines (Sulzer et al., 2000) (and personal communication). Further, over-expression of VMAT2 in PC12 cells lessens the toxic effects of cytosolic dopamine; reserpine has the opposite effect and potentiates dopamine’s toxicity (Weingarten and Zhou, 2001). Primary cultures from VMAT2 knockout mice are also more sensitive to the toxic effects of increased dopamine synthesis (Kariya et al., 2005) and decreased levels of VMAT2 can potentiate the neurotoxic effects of amphetamines (Larsen et al., 2002). In vivo evidence for the importance of VMAT2 in regulating cytosolic dopamine has come from studies in both mice and flies. VMAT2 knockout mice are more sensitive to neurodegenerative effects of amphetamine (Fumagalli et al., 1999). Similarly, reducing expression of the fly isoform of VMAT potentiates the death of dopaminergic neurons in a fly model of Parkinson’s disease (Sang et al., 2007) and VMAT2 knockdown mice show an increase in dopaminergic cell death (Caudle et al., 2007). Conversely, overexpression of Drosophila VMAT exerts a neuroprotective effect on dopaminergic neurons in the fly (Sang et al., 2007).

11

Behavioral Genetics of Animal Models

One of the most interesting aspects of vesicular transporters from a neuroscience perspective is the contribution that they may have to the function of the nervous system in the intact organism, and their impact on behavior. Drugs that influence the function of VMAT such as reserpine can have profound central and peripheral affects (Freis, 1954; Beers and Passman, 1990) suggesting that altered expression might also have important behavioral sequelae. Knockout and transgenic technologies in rodents and other experimental animals have allowed this to be directly tested for most of the known vesicular transporters.

11.1 Mouse Models The first vesicular transporter gene to be knocked out in mice was VMAT2 (Fon et al., 1997; Takahashi et al., 1997b; Wang et al., 1997). Since then, additional knockouts have been generated for VGAT (Wojcik et al., 2006), VGLUT1 (Fremeau Jr et al., 2004a; Wojcik et al., 2004) and VGLUT2 (Moechars et al., 2006). The available knockouts are generally lethal as homozygotes but important behavioral information has been obtained from studying the heterozygotes. In addition, VMAT2 (Mooslehner et al., 2001) and VAChT (Prado et al., 2006) knockdown animals have been generated. These animals survive through adulthood and have been subjected to extensive phenotypic analyse. VMAT2 heterozygotes show an increase in locomotor behavior in response to the aminergic drugs apomorphine, cocaine, and amphetamine, suggesting that the synaptic response to amine release has been increased (Fon et al., 1997; Takahashi et al., 1997b; Wang et al., 1997). However, these mice do not appear to undergo sensitization in response to chronic cocaine administration (Fon et al., 1997; Wang et al., 1997). That is, they do not show a potentiated behavioral response to cocaine following repeated administration of this drug. Rather, the increase in locomotion observed after the first cocaine treatment is equivalent to the response to later treatment. VMAT2 heterozygotes also show a reduced preference for amphetamine in a place preference test (Takahashi et al., 1997b). Both sensitization and place preference tests are important models for the addictive properties of drugs of abuse. In addition, sensitization may reflect an alteration in the synaptic signaling pathways relevant to the long-term changes that underlie addiction. Lack of sensitization in VMAT2 heterozygotes suggest that these mice are not capable of mounting a similar adaptive response, and that the relevant aminergic signaling pathways may be significantly altered. Heterozygous VMAT2 mice also show an increased locomotor response to ethanol (Wang et al., 1997). Moreover, males may show an increase in ethanol consumption (Hall et al., 2003). VMAT2 knockdown mice show a downregulation of substance P expression and an upregulation of enkephalin

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expression, suggesting potential effects on signaling pathways mediated by endogenous opioids in the striatum (Colebrooke et al., 2007). Together, this data shows that multiple signaling pathways are altered in response to a decrease in VMAT2 expression - all of which may contribute to the observed behavioral changes. The importance of VMAT2 expression for complex behaviors other than those associated with drugs of abuse are underscored by a report showing that VMAT2 heterozygotes perform worse than wild type littermates in several rodent models of clinical depression (Fukui et al., 2007). This is not surprising given the profound behavioral deficits seen in animals and humans treated with the VMAT inhibitor reserpine (Freis, 1954). However, in contrast to early pharmacological studies, the increase in depressive like behaviors in VMAT2 heterozygotes is clearly associated with the specific loss of VMAT2, rather than inhibition of VMAT1, which could cause fatigue due to cardiovascular effects. VAChT knockdown rather than knockout mice have been generated and subjected to both biochemical and behavioral analyses (Prado et al., 2006). In the central nervous system, extracellular acetylcholine is decreased by 30% in the VAChT knockdown mice, and KCl-induced release of acetylcholine also is reduced. Surprisingly, total tissue levels of acetylcholine were increased in both heterozygotes and homozygotes relative to wild type (Prado et al., 2006). The reason why extracellular acetylcholine would be decreased while total tissue acetylcholine is increased has not yet been resolved. However, it is possible that the elevation in tissue content results from an increase in the cytoplasmic pool of acetylcholine that occurs because of reduced transport into vesicles. In support of this interpretation, blockade of VAChT by vesamicol during prolonged stimulation of the NMJ decreases release, but increases tissue content (Collier et al., 1986). Similarly, VAChT mutants in C. elegans (unc-17) show an increase in total tissue acetylcholine (Hosono et al., 1987). VAChT knockdown mice survive but have neuromuscular defects prohibiting the analysis of complex behaviors (Prado et al., 2006). The neuromuscular function of VAChT heterozygotes minimally differs from wild type (Prado et al., 2006). However, several interesting, and relatively circumscribed behavioral deficits were observed in the heterozygous VAChT knockdown mice (Prado et al., 2006). Learning and memory for an avoidance task that depends on hippocampal and amygdalal networks was normal; in contrast showed the heterozygotes, a defect in remembering novel versus familiar objects. Heterozygotes also showed less habituation to an unfamiliar intruder, as if they had difficulty in learning to recognize novel versus familiar animals as well as objects. This apparent defect in social memory could be rescued with drugs that increase cholinergic tone, indicating that the behavioral phenotype was not the result of a developmental defect, but rather a decrease in acetylcholine release. In a clinical settings, the symptomatology of dementia is associated with decreased cholinergic tone, and drugs that increase cholinergic signaling have a mild but significant effect on prolonging cognitive function (Ringman and Cummings, 2006). The VAChT knockdown mice may be useful to test additional strategies for the treatment of cognitive symptoms caused by decreased acetylcholine release. VGLUT1 homozygous knockout pups feed poorly and show progressive, lethal neurological defects preventing additional studies of more complex behaviors (Fremeau Jr et al., 2004a; Wojcik et al., 2004). Interestingly, the onset of neurological defects appears to coincide with the developmental switch from VGLUT2 to VGLUT1 as the predominant CNS isoform. In contrast, heterozygotes survive through adulthood allowing behavioral studies. The VGLUT1 heterozygotes show a decrease in an exploratory paradigm often used as an animal model of anxiety (Tordera et al., 2007). In addition, these animals show an increase in immobility in the forced swim test, a model of depressive like behaviors in rodents. The precise relationship of VGLUT1 expression to anxiety and mood states remains unclear; nonetheless, these data show that, similar to VMAT2 and VAChT, relatively modest reductions in VGLUT1 expression can have profound effects on complex behaviors. VGLUT1 heterozygotes also show a decrease in recognition of novel objects, but no change in spatial memory (Tordera et al., 2007). The preservation of spatial memory may reflect the dependence of this process on VGLUT2. VGLUT2 knockout homozygotes die at or immediately after birth, presumably as result of respiratory failure, since lung alveoli appear to be non-functional (Moechars et al., 2006). Since the pathways that express VGLUT2 in the medulla control respiratory rhythms, this phenotype was suggested to be the result of a central defect (Moechars et al., 2006). Electrophysiological studies revealed that evoked glutamatergic

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responses were decreased in thalamic neurons but not in the hippocampus (Moechars et al., 2006). Preservation of VGLUT2-mediated hippocampal glutamate release may explain the apparent preservation of spatial memory in VGLUT1 heterozygotes (see above). Unlike VGLUT1 heterozygotes, VGLUT2 heterozygotes do not show defects in learning or memory formation, and are surprisingly similar to wild type animals in several other behavioral tests, including their acute response to painful stimuli (Moechars et al., 2006). However, they show a surprising reduction in the sensation of neuropathic pain. In these experiments, two roots of the sciatic nerve were severed, an established procedure that usually causes an increased sensitivity to relatively innocuous stimuli distal to the lesion. Unlike wild type controls, the VGLUT2 heterozygotes did not show a heightened response to either mechanical stimulation or cold after the nerve roots were severed. In wild type animals, this response requires glutamatergic circuits in the thalamus, and the loss of VGLUT2 may interrupt the flow of sensory information in the thalamus and/or the remodeling of thalamic circuitry. Additional, relatively subtle defects in the VGLUT2 heterozygotes reinforce the notion that thalamic function is impaired. These include a rapid extinction of conditioned taste aversion, similar to the effects of thalamic lesions in the rat. Further experiments will be needed to determine whether or not the thalamus is the true locus of the behavioral defects seen in the VGLUT2 heterozygotes. Nonetheless, these findings suggest that VGLUT2 could be a valuable and unexpected target for the treatment of neuropathic pain.

11.2 C. elegans As described above, behavioral genetic studies of C. elegans were critical to the initial identification of VAChT (Alfonso et al., 1993), VGAT (McIntire, 1993; McIntire et al., 1997) and VGLUT1 (Avery, 1993; Bellocchio et al., 1998). The behavioral phenotypes of the three C. elegans mutants unc-17 (VAChT), unc-47 (VGAT), and eat-4 (VGLUT) were known long before they were determined to encode vesicular transporters (Brenner, 1974; Alfonso et al., 1993). Both unc-17 and 47 were identified as uncoordinated mutants in the original behavioral screens in this system by Sydney Brenner and colleagues (Brenner, 1974). Interestingly, the unc-17 mutant was shown to be resistant to a cholinesterase inhibitor in the original 1974 report; it would take nearly 20 years to determine the relationship of the gene to cholinergic signaling (Alfonso et al., 1993). Eat-4 was identified as a mutant defective in eating and found to show abnormal pharyngeal muscle movements (Avery, 1993). It was later shown to have severe defects in the normal withdrawal response to being touched on the ‘‘nose,’’ as well as defects in the response to odorants (Berger et al., 1998). The defect in withdrawal was shown to be the result of abnormal habituation; eat-4 worms show a wild-type response to a single tap, but habituate rapidly and unlike wild-type worms do not respond to additional taps (Rankin and Wicks, 2000). The C. elegans ortholog of VMAT is encoded by cat-1 (Duerr et al., 1999). Mutants show a defect in their ability to regulate locomotor behavior in response to food (Duerr et al., 1999). This phenotype could result from decreased signaling in several aminergic pathways known to control food intake in C. elegans. Dopaminergic mechanosensory neurons that sense the presence of bacteria provide information about food availability (Kindt et al., 2007) and exogenous application of octopamine will cause C. elegans to mimic behaviors seen during starvation (Horvitz et al., 1982). Serotonergic circuits regulate food-induced slowing, and navigation during foraging (Dernovici et al., 2007) as well as the function of NMJ (Nurrish et al., 1999). Although some signaling pathways regulating these responses have been investigated, they are not completely understood (Suo et al., 2006; Dernovici et al., 2007; Kindt et al., 2007). The cat-1 mutants may be useful to further dissect molecular mechanisms underlying the response of C. elegans to food, and the potential contribution from presynaptic elements including VMAT.

11.3 Drosophila In Drosophila, mutations in both VAChT (Kitamoto et al., 2000) and VGLUT (Daniels et al., 2006) have been reported. Drosophila VGLUT (dVGLUT) mutants die as larva and electrophysiological analyses have been used to analyze the number of transporters per SV (see above). However, the relatively short life span

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of the dVGLUT homozygotes prevents an in depth behavioral analysis and the phenotype of the heterozygotes has not yet been reported. Two mutant alleles of Drosophila VAChT have been generated, including VAChT1 which is embryonic lethal, and VAChT2 (Kitamoto et al., 2000). VAChT2 larvae can survive through the second stage (instar) of development and locomote more slowly than wild type animals (Kitamoto et al., 2000). Similar to VAChT knock-down mice, heterozygous dVAChT adults can survive. Although the behavior of adult heterozygotes has not been reported, electrophysiological assays suggest that during periods of sustained vesicle release, at least one circuit in the adult CNS fails to maintain normal levels of acetylcholine release (Kitamoto et al., 2000). It is useful to compare this phenotype to electrophysiological results at the frog NMJ. In the frog, quantal size is decreased under periods of sustained release, an effect that may relate to the failure of the VAChT to adequately fill SVs (Naves and Van der Kloot, 1996). Similarly, it is possible that in the VAChT2 fly mutant, lower transport activity prevents SVs from filling with acetylcholine when they are being rapidly released and recycled. Current molecular genetic techniques in Drosophila are particularly well suited to in vivo overexpression studies, making it possible to determine how increased expression of a vesicular transporter may affect behavior. Over-expression of Drosophila VGLUT is larval lethal, but only electrophysiological experiments have been reported. In contrast, flies over-expressing DVMAT survive through adulthood (Chang et al., 2006). The widely used GAL4/UAS system (Brand and Perrimon, 1993) allows promoters and expression transgenes to be mixed and matched, and in the DVMAT experiments, the Dopa decarboxylase (Ddc) promoter was used to drive expression in serotonergic and dopaminergic cells (Li et al., 2000). The resulting changes in behavior, presumably due to an increase in dopamine and serotonin release, included an increase in motor and courtship behavior and a decreased behavioral response to cocaine (Chang et al., 2006). Importantly, these data indicates that a transporter–induced increase in quantal size, repeatedly demonstrated in vitro, can also have significant behavioral effects in an intact organism.

11.4 Zebrafish Over the past 20 years the zebrafish Danio rerio has emerged as an important model system. Mutation of the zebrafish VGLUT2a gene (the blu mutant) has been shown to have defects in spatial acuity, and in resolving rapidly changing visual stimuli, which decrease their ability to capture prey (Smear et al., 2007). VGLUT2a is partially responsible for glutamatergic transmission at the retinotectal synapse and during high-frequency stimulation, signaling at this site rapid fatigues in the mutant (Smear et al., 2007). Interestingly, an expansion of the retinotectal field was also detected in the mutant, indicating that partially compromising glutamate release can yield significant changes in neural circuitry, which is likely to contribute to the observed perceptual deficits. These results highlight the idea that some changes in behavior that result from altered vesicular transport may not be readily apparent from casual observation of the animal. Beyond the defects described above, VGLUT2a blu mutants are surprisingly intact. The ability of altered vesicular transporter to manifest as relatively subtle changes in behavior is relevant to their potential affects on neuropsychiatric illness in humans.

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Human Genetic Studies and Disease Models

12.1 Genetics Most neuropsychiatric illnesses are thought to be subject to complex mechanisms of inheritance that involve either multiple genes and/or interactions between genes and environmental factors. For plasma membrane neurotransmitter transporters, several polymorphisms may contribute the inheritance of clinically relevant traits, but only the mutation of NET is linked to the inheritance of a disease (orthostatic hypotension) (Hahn and Blakely, 2007). To date, polymorphisms in vesicular transporters have not been linked to any diseases or syndromes that are inherited in a strictly Mendelian fashion. However, polymorphisms may play a contributory role in some neuropsychiatric disorders.

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The possibility that changes in VMAT function may be relevant to psychiatric illness has prompted a search in humans for polymorphisms in both coding and non-coding regions of VMAT2 gene (Uhl et al., 2000; Glatt et al., 2001). In vitro transport assays indicate that only one coding polymorphism decreases the apparent affinity of VMAT2 for serotonin (Burman et al., 2004). Polymorphisms in the coding region of the VMAT2 have not been linked to Parkinson’s disease (Glatt et al., 2006a); however, polymorphisms in upstream regulatory domains have been shown to alter VMAT expression (Zars, 2000; Lin et al., 2005) and may be protective for alcoholism (Lin et al., 2005). In addition, VMAT2 promoter haplotypes that confer increased expression of VMAT2 in vitro may provide some protection against the development of Parkinson’s disease in women with high exposure to pesticides (Glatt et al., 2006b). To date, human genetic studies attempting to link VMAT2 polymorphisms with psychiatric illness have mostly yielded negative results (Persico et al., 1995; Kunugi et al., 2001) (but see (Gutierrez et al., 2007)). Similarly, polymorphisms associated with human disease have not yet been reported for VAChT (Harold et al., 2003), VGAT, or VGLUTs.

12.2 Altered Expression in Human Disease In addition to genetic studies of VMAT polymorphisms, a number of labs have assessed VMAT expression levels in patients suffering from a variety of neuropsychiatric illnesses including schizophrenia, depression, Tourette’s syndrome, and bipolar disorder (Meyer et al., 1999; Zubieta et al., 2000, 2001; Zucker et al., 2002a, b; Toren et al., 2005). Children with ADHD show a decrease in VMAT2 levels on platelets (Toren et al., 2005), and VMAT2 binding in the thalamus and ventral brainstem of bipolar patients is elevated (Zubieta et al., 2000, 2001). Patients with Tourette’s syndrome show an elevated level of VMAT2 expression in the ventral striatum (Albin et al., 2003). Consistent with this finding, striatal release of dopamine in response to amphetamines is increased in Tourette’s patients (Singer et al., 2002). Increased VMAT2 activity also has been detected in platelets of patients with schizophrenia and depression (Zucker et al., 2002a, b). The apparent lack of specificity in the relationship of VMAT2 activity to any particular psychiatric illness is somewhat concerning, but in aggregate, may suggest that high VMAT2 expression represents a general risk factor for disorders linked to aminergic circuits. It should be emphasized, however, that the potential risk associated with excess VMAT2 is likely to be relatively small, and at most, part of a more complex constellation of genetic and environmental factors. VMATs may also function as tissue markers in disease, and similar to other aminergic markers, VMAT2 expression is decreased in psychostimulant addiction (Chang et al., 2007). VMAT2 has also been proposed as a specific marker for tumors derived from gastric enterochrommafin-like cells (Eissele et al., 1999) and possibly pancreatic tumors derived from beta cells (Weihe and Eiden, 2000). Similarly, antibodies to VAChT have been suggested as a possible index for cellular degeneration in illnesses that affect cholinergic neurons such as amyotrophic lateral sclerosis (ALS) (Weihe and Eiden, 2000). In addition, VAChT may be downregulated in striatal interneurons in Huntington’s disease (Smith et al., 2006) (but see also (Suzuki et al., 2001)). In the case of VGLUT, the association of glutamatergic circuits with both neurological and psychiatric disease has led to a number of studies characterizing the response of VGLUTs to neuronal injury as well as psychotropic drugs (e.g., (Kang et al., 2005; Kim et al., 2005)) and VGLUT has been suggested as a marker to follow glutamatergic circuits in schizophrenia (Eastwood and Harrison, 2005).

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Conclusion

A great deal of information on the function of vesicular transporters has been obtained, much of this facilitated by their molecular cloning. However, many questions remain, and some of these are surprisingly basic. We anticipate that the further application of genetic, biophysical and cell biological techniques will determine the precise mechanism of transport, and how regulation of vesicular transport contributes to synaptic signaling and complex behavior.

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Acknowledgments The authors are funded by the NIMH (MH076900), NIEHS (ES015747) and the Hatos Center for Neuropharmacology. We thank Drs. Richard J. Reimer and Maarten E.A. Reith for their very helpful comments.

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Olfactory Neural Signaling from the Receptor to the Brain

K. Touhara

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 142

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Signaling in Olfactory Sensory Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 142 OR Gene and Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 142 Assaying Odorant Responsiveness of ORs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 144 OR Pharmacology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145 Signal Transduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 146

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Development of the Olfactory Sensory Neuron . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 148 Neurogenesis and OR Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 148 One Neuron-One Receptor Rule . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 149 Axon Convergence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 149 Zonal Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150

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Signaling to the Olfactory Bulb . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 Odor-Evoked Activity Pattern . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 OR Map . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153 Odorant Sensitivity and Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154

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Olfactory Neuronal Network in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154

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Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 155

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2009 Springer ScienceþBusiness Media, LLC.

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Abstract: The olfactory system is unique among the five sensory modalities in having both a wide range of sensor proteins, called olfactory receptors (ORs), and an extremely well-organized spatial neural projection pattern of sensory neurons. ORs expressed in olfactory neurons function as sensors of various odorants and comprise one of the largest multigene families in organisms from fish to primates. In addition, the expression of ORs, which is highly controlled by a one neuron-one OR rule, helps instruct the neural development of sensory neurons. This multi-role characteristic of ORs forms the molecular basis for a highly discriminative and sophisticated olfactory sensory system. In the last decade or so, it has become increasingly clear how neural signals of olfactory sensory neurons are transmitted and how odorant identity is represented as an odor map in the olfactory bulb and in the brain. Our current knowledge of the structure and function of ORs from the standpoints of molecular biology, biochemistry, pharmacology, and neurobiology is reviewed, and also discuss how neural signaling in olfaction occurs between the receptor and the brain. Deciphering OR-mediated signaling should help us understand neural development and the plasticity of the olfactory sensory system. List of Abbreviations: CNG, cyclic nucleotide-gated; DG, 2-deoxyglucose; EG, eugenol; EOG, electroolfactography; GPCR, G protein-coupled receptors; GRK, G protein-coupled receptor kinase; MPOA-AH, medial preoptic area-anterior hypothalamus; OB, olfactory bulb; ORs, olfactory receptors; PKA, protein kinase A

1

Introduction

The olfactory system possesses remarkable discriminative power in distinguishing between thousands of different odorous chemicals present in nature. This is achieved by the expression of a vast repertoire of about 1,000 olfactory receptors (OR) in the olfactory sensory neurons of the nasal olfactory epithelium. The chemical information of odorants is converted to an electrical signal in the neurons, which is then transferred to the olfactory bulb (OB), the first relay center of the olfactory system. The pattern of OR activation elicited by the electrical signal in OB is then sent to the higher cortical areas of the brain where perception and identification of the odorant is established. Thus, ORs play a critical role in the first step of odorant perception. In addition to their role as odorant sensors, ORs appear to be also involved in the construction of the beautiful neural network that constitutes the olfactory system. The one neuron-one OR rule ensures the convergence of olfactory sensory neurons expressing the same OR onto specific glomeruli during axon guidance, resulting in a distinctive neural circuitry that allows the representation of an activation pattern specific to each odorant in the OB. This circuitry is maintained throughout life by virtue of an ability of olfactory neurons to regenerate correctly. This chapter focuses on the molecular mechanisms underlying the neural signaling and plasticity of the olfactory system primarily with respect to ORs. First, our current knowledge of the structure and function of ORs will be reviewed, and then the odorant signal transduction mechanism in the olfactory sensory neurons will be described. Second, how olfactory sensory neurons develop and how the spatial neural network is topologically established will be discussed. Finally, recent progress in our understanding of the mechanism of odorant signal transmission to the OB and from the OB to the brain will be evaluated.

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Signaling in Olfactory Sensory Neurons

2.1 OR Gene and Structure Based on evidence for the involvement of G protein-mediated signaling and channels in olfactory signal transduction (Pace et al., 1985; Sklar et al., 1986; Nakamura and Gold, 1987; Breer et al., 1990; Dhallan et al., 1990), Buck and Axel hypothesized that receptor proteins for odorants are expressed in the olfactory epithelium and belong to a family of seven-transmembrane G protein-coupled receptors (GPCR) (Buck, 2004). With the advent of state-of-the-art molecular biology techniques, a multigene family encoding receptors for odorants was identified and named ORs or odorant receptors (Buck and Axel, 1991).

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OR proteins are classified as members of the GPCR superfamily because they possess certain of the structural features of GPCRs (Buck and Axel, 1991; Strader et al., 1994) and function in coupling and activating heterotrimeric G proteins (Krautwurst et al., 1998; Kajiya et al., 2001). Besides sharing several stretches of amino acid sequence homology (Zozulya et al., 2001; Zhang and Firestein, 2002), members of the OR family of proteins are characterized by their possession of seven hydrophobic transmembrane domains, a disufide bond between cysteines in their extracellular loops, and an N-terminal glycosylation site (> Figure 8‐1) (Katada et al., 2004). The conserved sequence motifs probably contribute to the proper folding of ORs in the plasma membrane, allowing them to bind to odorants and couple with appropriate G proteins. In contrast, the transmembrane regions form the odorant-binding pocket. Consequently, amino acid sequence homology in these regions is quite low as each OR must be able to distinguish between a wide variety of odorant molecules in the ligand recognition spectra (Singer et al., 1996; Pilpel and Lancet, 1999; Floriano et al., 2000, 2004; Singer, 2000; Man et al., 2004; Katada et al., 2005). Initially, the OR family was estimated to contain several hundred members in rat (Buck and Axel, 1991). In the last several years, whole genome sequencing has allowed a comprehensive analysis of the OR gene family, revealing details of OR genomic structure and distribution in various organisms (Mombaerts, 2004). The most recent data on vertebrate ORs are shown in > Figure 8‐1. In mammals, the OR repertoire comprises 800–1500 members. This is in contrast to fish that have only about 100 members (Niimura and

. Figure 8‐1 Structure and topology of a typical OR (upper). Red, Variable amino acids in the OR family; blue, conserved amino acids in the OR family. TM, transmembrane domain. Numbers of OR genes in various organisms (lower)

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Nei, 2005b). Thus, OR gene expansion would seem to have occurred when mammals moved from an aqueous to a terrestrial environment. In vertebrates, a significant portion of the OR gene family is made up of pseudogenes; hominoids have high pseudogene content (50%), whereas mouse and dog have lower contents of about 20% and 25–30%, respectively (Gilad et al., 2004; Niimura and Nei, 2005b; Quignon et al., 2005). The OR family has undergone and continues to undergo rapid molecular evolution through tandem gene duplication and pseudognization in each animal lineage. The fraction of pseudogenes in the order rodents, monkeys, and humans has increased during evolution, suggesting that the loss of olfactory function coincides with loss of functional OR genes. Indeed, in whale and dolphin, where the auditory system is dominant, 70–80% of OR genes appear to be pseudogenes (Y. Go, personal communication). Generally, OR genes are constituted of a single exon (1 kb) that contains an intronless coding region of 1 kb and upstream exons, whose number can vary between one and four (Lane et al., 2001; Young and Trask, 2002). OR genes form genomic clusters and are distributed widely on chromosomes. The distribution, however, is not even, and thus, almost half of mouse OR genes are located on chromosomes 2 and 7, which have orthologous relationships with chromosome 11 in humans (Niimura and Nei, 2005a). Further, OR genes are highly polymorphic as suggested for human leukocyte antigen-linked OR genes (Ehlers et al., 2000; Eklund et al., 2000) and dog OR genes (Tacher et al., 2005), possibly accounting for individual differences in odor perception and sometimes causing specific anosmia.

2.2 Assaying Odorant Responsiveness of ORs To assess OR function, olfactory neurons were first targeted as a site for OR expression (Zhao et al., 1998; Touhara et al., 1999), because they were predicted to possess the machinery for both OR expression and the transmission of odorant signals. Adenovirus-mediated gene transfer was used to overexpress ectopic ORs in the olfactory epithelium and the ligand responses were measured either by electro-olfactography (EOG) (Zhao et al., 1998) or by Ca2+ imaging of infected neurons (Touhara et al., 1999). In addition, a genetargeting approach was utilized to tag defined OR-expressing olfactory neurons with green fluorescent proteins with the aim of recording responses of the fluorescent cells to cognate odorants (Bozza et al., 2002; Grosmaitre et al., 2006). Both virus-mediated gene transfer and the gene-targeting approach, however, were tedious and time consuming, resulting in a search for a more convenient heterologous expression system. At first, it was not easy to express ORs functionally in heterologous cells because ORs appeared not to be efficiently translocated to the plasma membrane. Partial success was achieved by adding an N-terminal leader sequence from other GPCRs. This resulted in limited expression of functional ORs in the plasma membrane and in odorant-response recordings in a heterologous system, such as HEK293 cells (Krautwurst et al., 1998; Wetzel et al., 1999; Kajiya et al., 2001). In these cells, the co-expression of tagged-ORs and a promiscuous G protein, Ga5, led to Ca2+ responses when the cells were stimulated with the cognate ligand (> Figure 8‐2) (Krautwurst et al., 1998; Kajiya et al., 2001; Touhara et al., 2006). In the absence of Ga5 expression, the ORs activated endogenous Gas upon ligand stimulation, resulting in an increase in the cAMP level in various mammalian cell lines (> Figure 8‐2) (e.g., HEK293, COS-7, and CHO-K1 cells) (Kajiya et al., 2001; Katada et al., 2003). This approach was considerably facilitated by the implementation of a luciferase-reporter assay system using the zif268 promoter, which allowed luminescent detection of cAMP increases upon stimulation with an odorant (> Figure 8‐2) (Katada et al., 2003). Xenopus laevis oocyte is another good heterologous expression system for ORs (> Figure 8‐2) (Katada et al., 2003; Abaffy et al., 2006; Touhara et al., 2006). None of these expression systems, however, were applicable to all ORs. The possibility of chaperone involvement in membrane translocation of ORs was next investigated. The expression of the one transmembrane protein RTP1 was found to enhance cell surface expression of ORs and deorphanize certain ORs (Saito et al., 2004). In addition, Ric8B, a putative guanine nucleotide exchange factor for Gaolf, was recently found to promote efficient signal transduction by ORs (Von Dannecker et al., 2006). Thus, the current consensus is that the OR expression/translocation problem would be solved if all the factors required for OR expression in olfactory neurons, including those mentioned earlier, could be expressed

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. Figure 8‐2 Signaling cascades utilized to assay odorant responsiveness of an OR in heterologous cells. (Left, upper) Eugenol-induced Ca2+ responses in HEK293 cells coexpressing mOR-EG and Ga15. (Left, lower) Electrophysiological recording of eugenol-induced currents in X. laevis oocytes expressing mOR-EG and either Ga15 or CFTR (cystic fibrosis transmembrane regulator). (Right, lower) Eugenol responses to mOR-EG detected by zif268-promoter-mediated expression of luciferase activity in PC12h cells. (Right, upper) Eugenol-induced cAMP increases in mOR-EG-expressing HEK293 cells (see Katada et al., 2003, and Touhara et al., 2006, for details)

in heterologous expression systems. It would also be helpful to develop methodologies that maintain ORs stably on the cell surface, because ORs have been shown to undergo continuous internalization and recycling in heterologous cells (Jacquier et al., 2006).

2.3 OR Pharmacology Mouse mOR-EG (MOR174-9), the best-characterized OR so far, was originally isolated from a single eugenol (EG)-responsive neuron by Ca2+-imaging and single-cell RT-PCR techniques (Kajiya et al., 2001). Mouse mOR-EG recognizes at least 22 odorants that share certain molecular determinants with EC50 values ranging from a few micromolars to several hundred micromolars (Katada et al., 2005). Structure–activity relationship studies with mOR-EG have suggested that ORs have a broad but selective molecular receptive range, and that the selectivity is determined by the shape, size, and length of the ligand (> Figure 8‐3a). This pharmacological concept is a common feature of many ORs, as has been shown by studies on rat I7 (Araneda et al., 2000) and other deorphanized ORs (Wetzel et al., 1999; Bozza et al., 2002; Gaillard et al., 2002; Levasseur et al., 2003; Spehr et al., 2003; Matarazzo et al., 2005; Shirokova et al., 2005). Thus, odorant information about a specific odorant results from the combined activation of the array of ORs activated by the odorant (Malnic et al., 1999; Kajiya et al., 2001; Touhara, 2002). As is the case of other GPCRs, odorants can function both as agonists and antagonists of ORs, suggesting that the interactions between ORs and odorants is quite complicated. For example, the EG

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. Figure 8‐3 (a) Dose-dependent Ca2+ response profiles for various ligands in HEK293 cells expressing mOR-EG and Ga15. mOR-EG responds to eugenol, vanillin, and ethyl vanillin with different EC50 values. (b) A model for binding of eugenol to mOR-EG and the orientation of amino acids in the binding site. Eugenol is shown in light gray, and TMIII, V, and VI are shown as dark gray ribbons (modified from Katada et al., 2005)

response of mOR-EG was potently blocked by some structurally related odorants, such as methyl isoeugenol and isosafrol (Oka et al., 2004a, b). This antagonism was observed in olfactory neurons expressing mOR-EG in an intact olfactory epithelium slice preparation (Omura et al., 2003; Oka et al., 2004b). Further, quantitative recording using the Ca2+-imaging technique demonstrated that the antagonist response was partially inhibited in the OR-defined glomerulus (Oka and Touhara, unpublished observation). Thus, in complex mixtures of odorants, competition between the odorants for antagonism of the OR is likely to be another factor affecting the encoding of receptor information in the olfactory system. Multiple alignment analysis of the OR superfamily has revealed the existence of highly conserved and variable regions that are likely to be involved in structural organization and ligand recognition, respectively (Buck and Axel, 1991; Zhang and Firestein, 2002). A systematic experimental approach to decipher the odorant binding site was undertaken for mOR-EG (Katada et al., 2005). Functional analysis of site-directed mutants and ligand docking simulation studies were employed to define the odorant-binding environment. Most of the critical residues involved in odorant recognition were found to be hydrophobic and located within the binding pocket formed by transmembrane domains TM3, 5, and 6 (> Figure 8‐3b) (Katada et al., 2005). The spatial location of the binding pocket is similar to that of other biogenic GPCRs; however, in contrast to typical GPCRs where multiple electrostatic interactions with ligands are the rule, hydrophobic amino acids appear to be the rule for odorant recognition by ORs. Further, the accuracy of the binding model was validated by the fact that single amino acid changes caused predictable changes in agonist and antagonist specificity (Katada et al., 2005). Thus, ligand information appears to be transduced from the three-dimensional configuration that the binding pocket adopts when bound to a specific odorant ligand.

2.4 Signal Transduction The odorant signals of ORs are converted to electrical signals in olfactory sensory neurons (> Figure 8‐4a). The first step in signal transduction is the activation of an olfactory G protein, Gaolf, by the activated odorant-bound OR. Certain amino acids in the C-terminal domain and third intracellular loop of ORs appear to be involved in coupling and activating Gaolf (Katada et al., 2004) (Kato and Touhara, unpublished results). Unlike rhodopsin, signal amplification does not occur in the OR-G protein activation cycle (Bhandawat et al., 2005). Instead, Ric-8B, a guanine nucleotide-exchange factor, enhances accumulation of Gaolf at the cell membrane, which improves the efficiency of OR coupling to Gaolf (Von Dannecker et al., 2006). The activated Gaolf in turn stimulates adenylyl cyclase III, resulting in an

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. Figure 8‐4 Signaling in olfactory sensory neurons. (a) Odorant-induced signal transduction cascade. An olfactory receptor (OR), which is properly translocated to the plasmamembrane by virtue of a chaperone RTP, binds an odorant and couples to stimulatory G protein (Gaolf), which in turn activates type III adenylyl cyclase (ACIII), resulting in a cAMP increase followed by an opening of cyclic nucleotide-gated (CNG) channel and Cl channel. (b) Desensitization of olfactory neurons. The Ca2+–calmodulin (CaM) complex negatively regulates the CNG channel activity and inhibits ACIII via calmodulin-dependent kinase (CaMKII). An OR could be phosphorylated by G protein-coupled receptor kinase (GRK), resulting in returning to the inactive state

increase in the neuronal cell cAMP level. The elevated cAMP interacts with and opens up the cyclic nucleotide-gated (CNG) channel, leading to cation influx and depolarization of the receptor neuron. Also, Ca2+ activates a Cl channel, leading to an efflux of Cl, thereby further contributing to the amplification of sensory depolarization. Bestrophin-2 is a candidate for a molecular component of the olfactory Ca2+-activated Cl channel (Pifferi et al., 2006; Tsunenari et al., 2006). Gene knockout mice lacking either of the three molecular components (i.e., Gaolf, adenylyl cyclase III, or the CNG channel) failed to respond to odorants, suggesting that the cAMP cascade is dominant in transmitting the odorant signal in olfactory neurons (Brunet et al., 1996; Belluscio et al., 1998; Wong et al., 2000). However, it was later demonstrated that CNGA2 knockout mice could detect a subset of odorants, including 2-heptanone and 2, 5-dimethylpyrazine found in urine (Lin et al., 2004). Two additional olfactory pathways have been proposed: one involves membrane-bound guanylyl cyclase and phosphodiesterase type 2 (Fulle et al., 1995; Juilfs et al., 1997; Meyer et al., 2000), and the other involves phospholipase C and the TRP channel (Schild and Restrepo, 1998). The presence of multiple pathways in olfactory neurons may allow mice to respond readily to complex biological signals for social and sexual communication. The activated olfactory neurons must return to the steady state, which is referred to as desensitization (> Figure 8‐4b). As with other GPCRs, G protein-coupled ORs could be phosphorylated upon odorant binding by protein kinases, such as protein kinase A (PKA) and G protein-coupled receptor kinase (GRK), which would result in desensitization (Dawson et al., 1993; Schleicher et al., 1993; Lefkowitz, 2004). Indeed, knockout mice lacking GRK showed slower recovery from cAMP increases, implicating the role of GRK in OR desensitization (Peppel et al., 1997). In addition, it was shown that PKA-mediated phosphorylation and subsequent internalization of ORs along with b-arrestin leads to reduced amounts of ORs in the membrane (Mashukova et al., 2006). Although direct evidence for OR phosphorylation has not been obtained, desensitization mechanisms are likely to exist at the level of the OR. In addition, elevated [Ca2+]

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via influx through the CNG channel leads to the closing of the CNG channel, constituting a feedback mechanism for Ca2+ influx (Kurahashi and Shibuya, 1990; Zufall et al., 1991; Kurahashi and Menini, 1997). Ca2+ also negatively regulates adenylyl cyclase activity via calmodulin-dependent kinase (Wei et al., 1998). These Ca2+-mediated negative feedback mechanisms would allow activated olfactory neurons to go back to the steady state and prepare for the next stimulus.

3

Development of the Olfactory Sensory Neuron

3.1 Neurogenesis and OR Expression At approximately mouse embryonic day 10 (E10), a thickening of a region of the frontonasal surface epithelium leads to the formation of the olfactory placode that then starts to generate immature neurons (Balmer and LaMantia, 2005; Beites et al., 2005). At E11, the olfactory primordia are visible, and neural adhesion molecule (NCAM)-labeled neurons can be seen in the placode and nascent olfactory nerve. The onset of OR expression was determined at E12–14 by using in situ probes for several ORs (Strotmann et al., 1995; Sullivan et al., 1995). Recently, using specific antibodies against ORs, OR proteins were visualized in dendritic knobs of olfactory sensory neurons as early as E12 before the initiation of ciliogenesis (Barnea et al., 2004; Strotmann et al., 2004; Schwarzenbacher et al., 2005). After the formation of olfactory cilia at E13, OR proteins appear to migrate into the developing cilia, and strong OR antibody labeling is detectable in the expanded tips of the clia. Up to E18, the continued increase in the number of cilia per neuron is accompanied by concomitant increases in OR immunostaining. Microarray analysis of the developmental course of OR gene expression has revealed that the number of transcribed OR genes remains relatively low until birth (100–200 ORs). After birth, this number increases dramatically to be detected only after birth and reaches a peak at 2–3 weeks postnatal time (500 ORs) (Zhang et al., 2004). Unlike other general neurons, olfactory sensory neurons are regenerated throughout life. Neural stem cells in the olfactory epithelium called globose basal cells give rise to transit amplifying progenitors that express Mash1 (> Figure 8‐5). This is followed by immediate neuronal precursors expressing Ngn1, which eventually become NCAM-positive immature olfactory neurons (Schwob, 2002). Globose basal cells may

. Figure 8‐5 A model of the olfactory neuronal development from stem cells called globose basal cell (GBC). The molecular markers for each stage are shown: Mash1, mammalian achaete scute homolog 1; Ngn1, nuerogenin1; NCAM, neural cell adhesion molecule; OMP, olfactory marker protein

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also generate sustentacular cells to support epithelium structure and olfactory ensheathing cells for the envelopment of the olfactory nerve (Huard et al., 1998). OR expression begins at the immature neuron stage, wherein OR-mediated Gas-cAMP signaling ensures precise axonal projection and convergence of olfactory neurons in glomeruli of the OB (Imai et al., 2006). In mature neurons that have accomplished formation of the neural network, Gas expression is replaced by that of Gaolf, which plays a role in odorant signal tranduction. The temporal and spatial development of olfactory neurons appears to be carefully controlled by the orchestrated expression of a wide range of genes.

3.2 One Neuron-One Receptor Rule Each olfactory sensory neuron expresses just one OR out of a repertoire of about a thousand (Serizawa et al., 2004; Shykind, 2005). The one neuron-one receptor rule has been confirmed by a variety of techniques, including in situ hybridization (Ngai et al., 1993; Ressler et al., 1993; Vassar et al., 1993), single cell RT-PCR analysis (Malnic et al., 1999; Touhara et al., 1999; Oka et al., 2006), and transgenic experiments (Qasba and Reed, 1998; Serizawa et al., 2000, 2003; Vassalli et al., 2002). Further, the OR appears to be transcribed from only one of the two alleles, either the maternal or the paternal allele (Chess et al., 1994; Ishii et al., 2001). This mutually exclusive expression pattern is preserved even between transgenes and endogeous copies, which is in support of the stochastic model (Qasba and Reed, 1998; Serizawa et al., 2000, 2003; Vassalli et al., 2002). Indeed, the single OR choice was observed by using a minigene containing only a few kilobases of DNA from the upstream region of the transcription start site, suggesting an important role for cis-regulatory elements in this process (Qasba and Reed, 1998; Vassalli et al., 2002; Oka et al., 2006). In contrast, the expression of a cluster of OR genes, including MOR28, in transgenic mice required a long upstream region of 100 kb containing an enhancer element referred to as the H-region (Serizawa et al., 2000, 2003). This locus control region, H, appears to associate with multiple OR promoters on different chromosomes and to act as a trans-acting enhancer element (> Figure 8‐6). The expression levels of ORs differ considerably by up to 300-fold as a result of unequal numbers of expressing cells and unequal transcript levels per cell (Young et al., 2003; Zhang et al., 2004). In this regard, it would be of particular interest if the incidence of expression correlated with the accessibility of the trans-regulating element, H, to each OR gene. In addition to positive regulation of OR gene expression by trans- and cis-regulatory elements, negative-feedback regulatory loops must also exist to suppress the expression of additional receptors (Serizawa et al., 2003; Lewcock and Reed, 2004; Shykind et al., 2004). It appears that functional OR proteins rather than OR mRNA play an inhibitory role in preventing further activation of other OR genes, although the molecular mechanism remains to be elucidated.

3.3 Axon Convergence Olfactory sensory neurons expressing a given OR project their axons to defined glomeruli in the OB (> Figure 8‐7) (Ressler et al., 1994; Vassar et al., 1994; Mombaerts et al., 1996). Genetic manipulation that results in either deletion or receptor swap suggests that the OR protein itself is instructive for the axon guidance of olfactory neurons (Wang et al., 1998). Indeed, OR proteins are present in olfactory axonal terminals (Barnea et al., 2004; Strotmann et al., 2004) as well as in cells of the cribriform mesenchyme involved in the fasciculation and sorting of outgrowing axons (Schwarzenbacher et al., 2006). A point mutation in an OR protein resulted in formation of novel glomeruli, indicating that the OR amino acid sequence encodes information about axonal identity (Feinstein and Mombaerts, 2004). The OR, however, is not the sole player in axonal convergence; membrane receptors such as the ephrin receptor and Kirrel2/3 could be coordinately involved in the determination of the sensory map in the OB (Cutforth et al., 2003; Serizawa et al., 2006). In addition, instructive axon guidance is not restricted to ORs as the b2-adrenergic receptor can substitute for an OR in glomerular formation, suggesting that GPCRs may share common functions with ORs (Feinstein et al., 2004). However, in this respect, it should be noted that both ORs and

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. Figure 8‐6 A model for mechanisms underlying one neuron-one OR expression. The locus control region (LCR) containing an enhancer element associates with multiple OR promoters on different chromosomes to activate OR gene expression. The expressed functional OR proteins inhibit expression of other ORs, resulting in one OR in a single neuron. Two distinct ORs expressed in different olfactory neurons are stained by different colors

the b2-adrenergic receptor couple to stimulatory G proteins, Gas. Consistently, it was found that Gas signaling is one of the positional determinants for the convergence of olfactory neurons onto specific glomeruli in the OB (Imai et al., 2006). Thus, OR proteins expressed on the olfactory neuronal cilia and dendrites function to recognize odorants from the external environment, whereas the same OR proteins localized on axon termini appear to function as critical factors for the transmission of information for proper axon guidance.

3.4 Zonal Expression In mouse and rat, in situ hybridization experiments using different OR probes suggested that each OR gene is expressed only in a certain region of the olfactory epithelium (> Figure 8‐7) (Nef et al., 1992; Strotmann et al., 1992, 1994; Ressler et al., 1993; Vassar et al., 1993). On this basis, the epithelium was divided into at least four spatial zones depending on the set of OR genes expressed there (Sullivan et al., 1996). The distinct spatial expression pattern appears to be conserved among vertebrate species, although the expression of fish OR was not as clearly segregated as that of mammalian OR (Ngai et al., 1993; Weth et al., 1996). Initially, the spatial zones observed in rodents were thought to be sharply segregated, but careful in situ hybridization studies revealed that each OR gene was expressed in a distinct zone in a continuous and overlapping manner (Norlin et al., 2001; Iwema et al., 2004; Miyamichi et al., 2005). Therefore, multiple overlapping bands are formed along the dorsomedial/ventrolateral axis, except that the boundary of the most dorsal region, corresponding to that of OCAM-positive neuronal layer, is maintained. There are exceptions to the zonal expression rule. First, some OR genes are expressed in the septal organ that is an isolated island of olfactory sensory neurons located on the nasal septum near the entrance of the nasopharynx (Kaluza et al., 2004; Tian and Ma, 2004). Second, the OR37 subfamily showed a patch-like distribution in clustered populations of neurons in central turbinates (Strotmann et al., 1999). Although

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. Figure 8‐7 Anatomy of the olfactory system and spatial distribution of olfactory neurons expressing a certain OR. Analysis of gene-targeted mice expressing LacZ in mOR-EG-expressing olfactory neurons shows that mOR-EG is expressed in the most dorsal epithelium (zone 1) and the neurons expressing mOR-EG send their axons to the specific glomeruli in the olfactory bulb

there are some promoter motifs that might restrict expression to specific domains in the nasal cavity, the regulatory factors that determine this positional information remain to be identified. Taken together, these results indicate that ORs play a critical role in implementing the proper spatial and temporal organization of the neuronal network during the development of olfactory sensory neurons.

4

Signaling to the Olfactory Bulb

4.1 Odor-Evoked Activity Pattern The OB is the first part of the brain to relay the neural signals of olfactory sensory neurons to secondary neurons, called mitral/tufted cells, which in turn project their axons into the central olfactory system (Mori et al., 1999; Shepherd et al., 2004). The most outer layer of the OB comprises approximately 1,800 glomeruli, in which the axons of olfactory sensory neurons form synapses with the dendrites of mitral/ tufted cells. Approximately 10,000 olfactory sensory neurons expressing the same OR send their axons to a few topologically fixed glomeruli, resulting in a pattern of activated glomeruli that reflects the original

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pattern of activated ORs in the olfactory epithelium (> Figure 8‐7) (Ressler et al., 1994; Vassar et al., 1994; Mombaerts et al., 1996). In other words, individual odorants activate distinct subsets of ORs, resulting in a glomerular activation pattern that is unique for each odorant in a stereotyped region of the OB called the ‘‘odor map’’ (Xu et al., 2000; Leon and Johnson, 2003; Mori et al., 2006). The odor maps in the OB can be assessed using several imaging methods such as Ca2+ imaging (Friedrich and Korsching, 1997; Wachowiak and Cohen, 2001; Oka et al., 2006), intrinsic imaging (Rubin and Katz, 1999; Uchida et al., 2000), and functional MRI (Xu et al., 2003), as well as by methods employing various neuronal activity markers such as 2-deoxyglucose (DG) and immediate early gene expression (see references in Shepherd et al., 2004). The 2-DG and gene expression methods allow the mapping of responses in the entire OB, but only with a single odorant per animal. In contrast, the Ca2+ and intrinsic imaging techniques allow recording of glomerular responses to different odorants at the level of a single glomerulus in the same animal, but imaging is limited to the dorsal and postero-lateral surfaces of the OB. Although functional MRI solves the inherent disadvantages of these two methods, the resolution of this method is not high enough to measure responses at the glomerulus level. Ca2+ imaging directly measures the Ca2+ influx into axon termini of olfactory sensory neurons, while intrinsic imaging picks up optical signals of hemoglobin in blood around glomeruli. Therefore, Ca2+ imaging is so far the best approach for performing response analysis for odor mapping in the OB. > Figure 8‐8 shows examples of glomerular responses to various odorants as detected by Ca2+ imaging in the dorsal OB. Different odorants elicit different glomerular activity patterns, but structurally related odorants activate similar sets of glomeruli, reflecting the fact that similar odorants are recognized by similar sets of ORs in the olfactory epithelium. Even for the same odorant, different concentrations elicit different patterns of activation in that more glomeruli are recruited at higher concentrations of odorants (> Figure 8‐8a) (Oka et al., 2006). There are many odorants that do not activate glomeruli in the dorsal region. For example, volatile odorants in urine do not elicit responses in the dorsal glomeruli, but evoke responses in the most ventral area as shown by monitoring immediate early gene expression (Schaefer et al.,

. Figure 8‐8 (a) Ca2+ responses of glomeruli to different concentrations of eugenol in the dorsal regions of the mouse olfactory bulb. (b) Identification of OR genes expressed in olfactory neurons innervating odorant-activated glomeruli. ORs shown on the bottom were identified by in vivo Ca2+ imaging followed by retrograde dye labeling and single-cell RT-PCR (see Oka et al., 2006, for details)

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2001). There appears to be a functional odor map in the OB, and glomeruli with similar response properties are often located in spatial proximity (Mori et al., 2006).

4.2 OR Map To determine if the odor map in the OB truly corresponds to the receptor code in the olfactory epithelium, Ca2+-imaging techniques were applied to transgenic mice in which defined OR-expressing neurons were tagged with fluorescent probes. The glomerulus innervated by mOR-EG-expressing neurons showed responses to EG, a cognate ligand for mOR-EG (Oka et al., 2006) (> Figure 8‐9). Similarly, I7, M71, and . Figure 8‐9 Ca2+ responses to eugenol or vanillin in mOR-EG glomerulus and mOR-EG-expressing olfactory neurons. The responses to eugenol in the mOR-EG-glomerulus (left) or mOR-EG-expressing neurons (right) were fitted to a dose–response curve (see Oka et al., 2006, for details)

MOR23 glomeruli exhibited responses to their corresponding ligands, octanal, acetophenone, and lyral (Belluscio et al., 2002; Bozza et al., 2002; Grosmaitre et al., 2006). These studies provided evidence that the responsiveness of a glomerulus was a reflection of that of the OR expressed by the innervating sensory neurons. This type of study could eventually reveal the entire OR map in the OB, but the methodology remains tedious and time consuming. An alternative strategy to define the connection between the activated patterns of glomeruli and corresponding OR-expressing neurons is to identify the expressed OR by combining Ca2+ imaging, retrograde dye labeling, and single-cell RT-PCR. This approach has been successfully performed to isolate the four most sensitive ORs for EG and methyl isoeugenol, and one sensitive OR for isovaleric acid in the dorsal OB area (> Figure 8‐8b) (Oka et al., 2006). Spatial and functional mapping of these OR-defined glomeruli revealed that the positional relationship of glomeruli varies considerably between individual mice and even between the left and right OB in the same animal (Oka et al., 2006). These results suggest that precise bulbar OR maps may differ between individuals, making it impossible to acquire a true odor map without repeated examinations in many animals.

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4.3 Odorant Sensitivity and Specificity OR sensitivity to odorants in heterologous expression systems turns out to be relatively high in comparison with the odorant sensitivity of the olfactory system; the threshold concentrations for the best ligand for each OR range from a few micromolars to several hundred micromolars (Mombaerts, 2004). This is not simply a problem with the heterologous expression system as the sensitivity range of olfactory sensory neurons expressing the same OR is similar (Oka et al., 2004b, 2006). However, the odorant sensitivity of glomeruli innervated by olfactory neurons expressing a defined OR appears to be about 1,000-fold higher than that of peripheral neurons (> Figure 8‐9) (Oka et al., 2006). In addition, the specificity of in vivo odorant response in an OR-defined glomerulus was different from that suggested by in vitro OR pharmacology in a heterologous system (> Figure 8‐9) (Oka et al., 2006). The apparently higher odorant sensitivity and specificity in vivo seems to be a consequence of the olfactory mucus that provides a place for efficiently concentrating and carrying odorants to the receptor site. Nonetheless, an important caveat that needs to be considered is the possibility that ligand specificity of ORs obtained in heterologous OR expression systems may not always reflect the specificities observed in the OB under physiological conditions in vivo.

5

Olfactory Neuronal Network in the Brain

Mitral and tufted cells that synapse with olfactory sensory neurons in glomeruli project their axons into the olfactory cortex through the lateral olfactory tract (Shipley and Ennis, 1996). Classical neuroanatomical tracing studies have revealed that the regions receiving input from the OB, referred to as the primary olfactory cortex, include the anterior olfactory nucleus (AON), olfactory tubercle (Tu), piriform cortex (Pir), anterior cortical amygdaloid nucleus (ACo), posterolateral cortical amygdaloid nucleus (PLCo), and the lateral entorhinal cortex (LEnt) (> Figure 8‐10) (Prices, 1990). To elucidate how the signal from a single glomerulus is organized within these cortical structures at the cellular level, a transneuronal tracer, lectin, was expressed in olfactory sensory neurons expressing the same OR by the transgenic approach,

. Figure 8‐10 Olfactory connection to the brain. The lower picture is a fluorescent image showing that neural tracer Dil injected to the glomerular layer in the olfactory bulb (OB) are found in the anterior olfactory nucleus (AON) and piriform cortex (Pir). OE, olfactory epithelium; OB, olfactory bulb; AON, the anterior olfactory nucleus; Tu, olfactory tubercle; Pir, piriform cortex; ACo, anterior cortical amygdaloid nucleus; PLCo, posterolateral cortical amygdaloid nucleus; LEnt, the lateral entorhinal cortex

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which resulted in the labeling of higher-order neurons receiving signals from a particular OR (Zou et al., 2001). It turned out that cortical neurons originating from one OR type tended to be clustered and the distribution was different from OR to OR with partial overlapping. Further, using an intermediate early gene (c-Fos) as a neuronal activation marker, single odorants were shown to elicit distinct activation patterns that nonetheless partially overlapped in the piriform cortex (Zou et al., 2005). More recently, by examining the temporal subcellular localization patterns of the mRNA of another intermediate early gene (Arc) upon odorant stimulation, it was demonstrated that some cortical neurons require a combination of multiple receptor inputs for activation (Zou and Buck, 2006). These results suggest that although the spatial representation created in the OB is partly preserved in the olfactory cortex, it leads mostly to a distinctly distributed activation pattern. Each of the cortical subregions project information into various areas in the brain (Zelano and Sobel, 2005). One such region is the orbitofrontal cortex, a prominent site of olfactory processing. Functional imaging studies using fMRI or PET allowed identification of specific areas for olfactory processing upon odorant stimulation. Increased activity was observed in the medial gyrus and lateral orbitofrontal gyrus upon stimulation with pleasant and unpleasant odors, respectively (Anderson et al., 2003; de Araujo et al., 2005). Another region is the hypothalamus that plays a role in regulating reproductive behavior. By targeting trans-neuronal tracers to GnRH neurons in transgenic mice, the medial preoptic area-anterior hypothalamus (MPOA-AH), where many GnRH neurons are located, was shown to receive olfactory input from various cortical areas (Boehm et al., 2005). In addition, a virus-mediated approach successfully labeled the olfactory–hypothalamus pathway (Yoon et al., 2005). Both of these studies provide a hint as to the nature of the neural circuitry involved in odorant-induced mating behavior. In this regard, it is interesting to note that PET studies showed sex-specific patterns of hypothalamic activity in humans upon stimulation with androstadienone, a putative human pheromone (Savic et al., 2001, 2005). Neuroanatomical and genetic approaches, in combination with functional imaging approaches, have revealed neural plasticity for odorant identification and discrimination, and hard-wired circuits for innate reproductive behavior via the main olfactory system.

6

Conclusions

Since the discovery of the OR superfamily, considerable progress has been made in understanding the molecular mechanisms of this highly discriminative molecular recognition system. Ligand recognition by broadly tuned ORs and the accompanying unique neural circuitry directed by OR expression provides the molecular basis for our ability to detect and discriminate among about 1,000 different odors. This sophisticated neural sensory system is continuously regenerated throughout life, providing at the same time remarkable reproducibility and plasticity of olfactory sensations. An unforeseen outcome of research into olfactory function was the surprising discovery that ORs also play an important role in neural network construction. Moreover, it is also possible that ORs function as pheromone detectors to regulate innate behavior and as chemical sensors in non-olfactory tissues. These multiple roles of ORs may constitute one of the main forces for the conservation of the considerable diversity within the OR family during evolution. Interestingly, the number of functional ORs differs from animal to animal, and this distribution seems to reflect differences in the animal’s environment as well as the degree to which the animal relies on the olfactory function for intraspecies communication. In this regard, the degree of molecular diversity of the OR family in various organisms can tell us a lot about the evolutionary processes of development and the retardation of this sensory system in certain animals, such as human. However, there still remain a number of unresolved issues that need to be investigated, prominent among which are the olfactory mapping processes in the brain. The final goal should be to understand human olfactory perception at the molecular and systems level together with previous psychophysical literature. The difficulty, however, is that our understanding of olfaction obtained by using the mouse model may not easily be applicable to human beings. Technical approaches, such as genetic manipulation utilized in mice, are not possible in human studies. Furthermore, there seem to be large individual differences between humans depending on previous exposure to and experience of odorants in their environment. For these reasons, human olfactory

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research will be more challenging. Nonetheless, I believe that the study of the olfactory systems of various animals, such as fish, rodent, primate, and human, provides an excellent way to understand the highly complicated nature of neural plasticity in the brain.

Acknowledgments I would like to thank members of Touhara lab for providing data. This work was supported in part by grants from the Ministry of Education, Science, Sports, and Culture (MEXT), the Japan Society for the Promotion of Science (JSPS), and the Program for Promotion of Basic Research Activities for Innovative Biosciences (PROBRAIN) in Japan.

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Phosphorylation

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The Function of CaM Kinase II in Synaptic Plasticity and Spine Formation

K. Fukunaga . N. Shioda . E. Miyamoto

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164

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CaMKII Activation in Hippocampal LTP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164 Basic Structure and Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164 Frequency Detector of Ca2+ Oscillations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165 Activity in Hippocampal LTP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165

3 3.1 3.2

CaMKII in Explicit and Implicit Memories . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167 Explicit Memory in Transgenic Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167 Implicit Memory in Transgenic Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167

4 4.1 4.2 4.3 4.3.1

CaMKII in Developmental Synaptic Plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 168 Interaction with NMDA Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 168 Interaction with MAGUK Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 Interaction with AMPA Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 Stargazin in GluR1 Delivery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 170

5 5.1 5.2 5.3 5.4

Phosphorylation-Dependent Synaptic Plasticity in Mature Synapses . . . . . . . . . . . . . . . . . . . . . . . . . 170 Molecular Switch of LTP and LTD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 170 Phosphorylation of GluR1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 170 Dephosphorylation of GluR1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171 PKC in Cerebellar LTD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 171

6 6.1 6.2

LTP Maintenance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172 Protein Synthesis-dependent LTP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172 Delivery of mRNA to Dendrite . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 173

7 7.1 7.2 7.3 7.4 7.5 7.6 7.7

Morphological Change in Spine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 173 Spine Formation in LTP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 173 Actin Reorganization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 173 PAK Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174 Ca2+ Mobilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174 Guanine-Nucleotide Exchange Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175 SynGAP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 176 EphB Receptor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 176

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Abnormal Spine Morphogenesis in Mental Retardation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 176 Genes Associated with Mental Retardation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 176 CaMKII in Angelman and ATR-X Syndromes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 177

9

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 177

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Abstract: Recent advances in molecular genetics provide strong evidence for a relationship between hippocampal long-term potentiation (LTP) and hippocampus-dependent memory. Genetic deletion of a crucial Ca2+/calmodulin-dependent protein kinase II (CaMKII) subunit or inhibition of its autophosphorylation blocks LTP induction and causes severe hippocampus-dependent memory deficits. Synaptic activity-dependent phosphorylation and trafficking of the GluR1 subunit through CaMKII activation account for hippocampal LTP. Moreover, CaMKII activation mediates rapid morphological changes in dendritic spines at excitatory synapses during LTP induction. In addition to the critical role played by CaMKII in synaptic plasticity in various brain regions, various psychotic disorders, including mental retardation, schizophrenia, and depression, alter CaMKII activity in specific brain regions correlated with changes in spine morphology. In this chapter, we focus on CaMKII-dependent mechanisms of memory formation and spine formation in the brain. List of Abbreviations: AMPA, (S)-a-amino-3-hydroxy-5-methyl-4-isoxazolepropionate; CaM, calmodulin; CaMKII, Ca2+/calmodulin-dependent protein kinase II; GluR, glutamate receptor; LTD, long-term depression; LTP, long-term potentiation; NMDA, N-methyl-D-aspartate; PKC, protein kinase C

1

Introduction

CaMKII is highly expressed in neurons where it can constitute up to 1–2% of total protein (Braun and Schulman, 1995). CaMKII functions in several Ca2+-mediated cellular processes, including neurotransmitter biosynthesis, neurotransmitter release, gene expression, cytoskeletal regulation, and neuronal plasticity (Colbran and Soderling, 1990; Hanson and Schulman, 1992b; Fukunaga and Miyamoto, 2000; Lisman et al., 2002; Hudman and Schulman 2002; Colbran 2004). Four CaMKII isoforms – a, b, g, and d – are each encoded by distinct genes in eukaryotes (Schulman and Hanson, 1993). These isoforms have sequence homologies of approximately 90% and 75% in the amino-terminal catalytic/regulatory domains (RGD) and carboxylterminal association domains, respectively (Tobimatsu and Fujisawa, 1989). The association domain is responsible for formation of a homo- or hetero-oligomeric complex containing 12 CaMKII subunits. Some CaMKII isoforms are targeted to specific cellular compartments: for example, the CaMKIIa isoform has been shown to bind to postsynaptic densities (Hanson and Schulman, 1992; Soderling et al., 2001), while the a, b and b0 isoforms bind to synaptic vesicles (Benfenati et al., 1992). The b isoform, but not the a isoform, binds F-actin in dendritic spines (Shen et al., 1998). Four isoforms, aB (Brocke et al., 1995), gA (Tobimatsu et al., 1988), gA0 (Takeuchi et al., 2002), and d3 (also termed the dB isoform; Edman and Schulman, 1992; Schworer et al., 1993; Mayer et al., 1994; Takeuchi et al., 1999) contain a nuclear localization signal (NLS) within the variable domain. Indeed, overexpressed aB and d3 isoforms are nuclear in neuroblastoma cells (Brocke et al., 1995), cardiac myocytes (Srinivasan et al., 1994), and NG108-15 cells (Takeuchi et al., 1999). We first focus on functional relevance of CaMKII isoforms to neuronal plasticity in memory functions.

2

CaMKII Activation in Hippocampal LTP

2.1 Basic Structure and Activation CaMKII is highly expressed in brain and enriched at synaptic structures, especially in postsynaptic densities. The enzyme forms a homo- or hetero-oligomeric structure composed of 12 subunits of the a or b isoform. CaMKII isoforms contain a highly conserved N-terminal catalytic domain (approximately 280 amino acids) followed by a 40 amino acid regulatory domain and a 150–220 amino acid C-terminus. Four CaMKII isoforms (a, b, g, and d) share 90% amino acid sequence identity in the catalytic domain and 75% amino acid sequence identity in the association domain. Variable domains exist between the regulatory and association domains and C-terminal of the association domain (> Figure 9-1). An interesting feature of the enzyme is that autophosphorylation of Thr-286 converts it from a Ca2+-dependent form to an -independent (constitutively active form) form (Miller et al., 1988; Schworer et al., 1988; Thiel et al., 1988; Lou and Schulman, 1989). The amino acid sequence of the RGD (residues 281–310) contains an

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. Figure 9-1 Structure and regulation of CaMKII. (a) CaMKII has an N-teminal catalytic domain, a central regulatory domain and a C-terminal association domain. The regulatory domain contains an autoinhibitory sequence and a calmodulin (CaM) binding site and exhibits several autophosphorylation sites including Thr286 and Thr305/ 306. Thr286 autophosphorylation leads to conformational changes allowing CaM binding, whereas Thr305/306 autophosphorylation inhibits CaM binding, thereby inhibiting Ca2+/CaM-dependent activity. (b) The sequence of regulatory domains (residues 281–310), which contains an autoinhibitory domain and a CaM binding domain. Residues 282–294 containing Thr286 interact with a hydrophobic pocket (T-site), whereas residues 297–300 occupy the substrate-binding site (S-site) of the catalytic domain as a pseudosubstrate. Thr286 autophosphorylation disrupts interaction with the T-site, thereby removing pseudosubstrate inhibition of the S-site by a conformational change. Binding of NR2B, an NMDA receptor subunit, to CaMKII also affects the T-site, thereby creating a constitutively active form of CaMKII without autophosphorylation. (c) CaMKII is formed rings of hexamers that stack on (the oligomer) each other via association domains. When Ca2+/CaM binds to one subunit of the holoenzyme, the subunit can phosphorylate an adjacent subunit in the presence of Ca2+/CaM (Hanson et al., 1994). This reaction requires Ca2+/CaM binding on both the ‘‘kinase’’ and ‘‘substrate’’ subunits and therefore requires two molecules of CaM to generate autonomous and trapped states. CaMKII homo- or hetero-oligomers are essential for persistent autophosphorylation, thereby constitutively active CaMKII

autoinhibitory and a CaM-binding domain. Residues 282–294 containing Thr-286 interact with the hydrophobic pocket (T-site), whereas residues 297–300 occupy the substrate-binding site (S-site) of the catalytic domain as a pseudosubstrate. Thr-286 autophosphorylation disrupts interaction with the T-site, thereby removing pseudosubstrate inhibition of the S-site via conformational change. Binding of NR2B, an NMDA receptor subunit, to CaMKII interacts with the T-site, thereby creating a constitutively active form of the protein in the absence of autophosphorylation (Bayer et al., 2001). Autophosphorylation itself is not a simple mechanism in CaMKII: when Ca2+/CaM binds to a subunit of the holoenzyme, the subunit can phosphorylate an adjacent subunit (Hanson et al., 1994) (> Figure 9-1). This reaction requires Ca2+/CaM binding on both the ‘‘kinase’’ and the ‘‘substrate’’ subunits and therefore requires two molecules of CaM to generate autonomous kinase activity and trapped states of CaM (Hanson et al., 1994). Thus, maximal activation of an autonomous kinase showing greater than 70% Ca2+-independence is obtained by

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autophosphorylation at 1–2 mol phosphate/mol holoenzyme. In addition, autophosphorylation of Thr-286 increases the dissociation constant for CaM from about 0.2 s for the unphosphorylated form to about 20 s for the phosphorylated form and results in trapping of CaM in the CaMKII molecule (Meyer et al., 1992). The multimeric form of the kinase facilitates autophosphorylation of Thr-286 and stabilizes the autophosphorylation state between subunits. Like autophosphorylation of Thr-286, positive cooperativity is also evident in CaM binding to each subunit. However, relatively high levels of Ca2+ (>1 mM) are required for high levels of CaM occupancy in CaMKII. Taken together, CaMKII autophosphorylation is associated with transient but significant increases in postsynaptic Ca2+ levels mediated by NMDA receptors following high frequency stimulation inducing LTP. A high threshold of Ca2+ is required to generate a positive cooperative state in CaMKII. Once CaMKII activity becomes autonomous (in the ‘‘on state’’), kinase activity can persist after intracellular Ca2+ concentrations return to basal levels below 100 nM. Thus autophosphorylation allows the enzyme to function as a memory molecule encoding the frequency of synaptic usage (Miller and Kennedy, 1986; Lisman, 1989).

2.2 Frequency Detector of Ca2+ Oscillations CaMKII function as a frequency detector of Ca2+ oscillations was experimentally demonstrated using an immobilization and superfusion system of a multimeric form of recombinant CaMKIIa and b. (De Koninck and Schulman, 1998). The affinity of CaMKIIb for CaM is greater than that of CaMKIIa. Generating autonomy depends on the frequency and duration of Ca2+ spikes as well as the free CaM concentration (De Koninck and Schulman, 1998). For example, a Ca2+ spike at 4 Hz with a 200 ms duration maximally generates autonomous kinase activity at different CaM concentrations. Interestingly, prephosphorylated CaMKII with 15% Ca2+-independence can further potentiate autonomous activity in response to a 1 Hz Ca2+ spike, which has no apparent effect on the unphosphorylated kinase. This effect is due to functional cooperativity described earlier and suggests that a highly autonomous kinase can maintain autophosphorylation at subthreshold activation (De Koninck and Schulman, 1998).

2.3 Activity in Hippocampal LTP In support to this idea, LTP-inducing high frequency stimuli leads to increased levels of Ca2+-independent CaMKII activity in hippocampal slices (Fukunaga et al., 1993; Fukunaga et al., 1995) (> Figure 9-2). In that system, the level of CaMKII Ca2+-independence was about 12% of the basal condition and increased to about 14% following LTP-inducing stimulation (> Figure 9-1). This increased autonomous kinase activity is relatively small but stable for at least 1 h. If LTP expression occurs in a limited dendritic spine, increases in observed autonomous kinase activity are apparently large enough to produce long-lasting biochemical changes in critical synapses. In contrast with the persistent activation of CaMKII, activation of CaMKIV and extracellular signal-regulated kinase (ERK) following hippocampal LTP is transient and returns to their basal activity within 30 min (Liu et al., 1999; Kasahara et al., 2001). Taken together, CaMKII autophosphorylation at Thr-286 is associated with synaptic changes underlying hippocampal LTP. By contrast, transient activation of CaMKIV and ERK is sufficient for triggering of gene expression associated with LTP maintenance (Kasahara et al., 2001; Sweatt, 2004). Following Thr-286 autophosphorylation, CaMKII autophosphorylation occurs at Thr-305 and/or Thr-306 (Colbran and Soderling, 1990; Patton et al., 1990), resulting in decreased affinity for Ca2+/CaM and a dramatic loss of Ca2+/CaM-dependent activity. Thus, Thr-305 and/or Thr-306 autophosphorylation are termed inhibitory autophosphorylation (Hashimoto et al., 1987; Kuret and Schulman, 1989). CaMKIIa autophosphorylation at Thr-305/306 in vitro reduces its affinity for the postsynaptic density (PSD) (Strack et al., 1997a, 1997b). Indeed, transgenic mice overexpressing forms of Thr-305/306 substituted with nonphosphorylatable amino acid residues show increased levels in PSD CaMKIIa and lowered thresholds for hippocampal LTP (Elgersma et al., 2002). Thus CaMKIIa association with the PSD modulates the threshold for LTP induction.

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. Figure 9-2 Persistent increases in constitutively active CaM kinase II activity contrast with transiently increased activity of CaM kinase IV in hippocampal LTP. (a) After high frequency stimulation (HFS) of afferents in the CA1 regions, the regions were dissected out at different time points and assessed for CaMKII and CaMKIV activities. (b) Electrophysiological recording illustrates the average changes in synaptic efficacy in the CA1 area of hippocampal slices produced by HFS. (c) Ca2+-independent activities of CaMKII were measured with syntide2 as substrate in CA1 areas of control (low frequency stimulation; LFS) slices (open squares) and slices receiving either high frequency stimulation (filled squares) or low frequency stimulation (open triangles) (Fukunaga et al., 1993). (d) CaMKIV activity in potentiated (HFS) or control (LFS) slices was measured with peptide-g as substrate in CA1 areas (Kasahara et al., 2001). Long-lasting elevation of CaMKII activity contrasts with a transient elevation in CaMKIV activity seen following hippocampal LTP

3

CaMKII in Explicit and Implicit Memories

3.1 Explicit Memory in Transgenic Mice CaMKIIa knockout mice provide strong evidence for function of the enzyme in LTP as well as in spatial learning, an explicit memory (Silva et al., 1992a; 1992b). Infection of hippocampal slices with vaccinia virus carrying a truncated, Ca2+-independent form of CaMKII also potentiates synaptic transmission and blocks LTP induction (Pettit et al., 1994). To further test the role of CaMKII autophosphorylation in LTP and memory, Mayford et al. (1995) conducted elegant studies with transgenic mice expressing CaMKIIAsp-286, which mimics autophosphorylation, in the hippocampus. In this case, the mutant kinase is 20to 30-fold more active in the absence of Ca2+ than the wild-type, and the CaMKII promoter was used to restrict transgene expression to the forebrain. Total Ca2+-independent activity increased from 7 to 14% in mutant mice without changing the total activity observed in adult hippocampus. This mutation is predicted

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to potentiate excitatory synaptic transmission and occlude LTP or undergo LTP more readily. However, synaptic transmission was not substantially altered: transgenic mice still exhibited normal LTP in response to stimulation at 100 Hz and in fact exhibited LTD in response to stimuli in the range of 1–10 Hz. It was also unanticipated that the size and direction of LTD produced following 5 Hz stimulation were strongly correlated with the level of Ca2+-independent CaMKII activity. Mayford and his colleagues also found that loss of hippocampal LTP in the range of the theta frequency was associated with selective impairment of spatial but not contextual memory, an observation consistent with the idea that theta rhythm is associated exploration in rodents (Bach et al., 1995; Mayford et al., 1995; Bejar et al., 2002). Furthermore, mutant mice expressing CaMKIIa-Asp-286 exhibited abnormal representation of place cells in the hippocampus during spatial learning (Rotenberg et al., 1996). To further address whether autophosphorylation of Thr-286 is critical for LTP induction in the hippocampus and spatial learning, an Ala-286 point mutation was introduced into the CaMKIIa gene, blocking autophosphorylation without affecting Ca2+/CaMdependent activity (Giese et al., 1998). Like CaMKII-Asp-286 mutants, Ala-286 mutants exhibit normal synaptic transmission. However, mutant mice showed no NMDA-receptor-dependent LTP in the hippocampal CA1 region and exhibited deficits in spatial learning in a Morris water maze. Consistent with deficits in spatial learning, the mice did not form stable place cells in the hippocampal CA1 region. Thus, Thr-286 autophosphorylation is a molecular switch underlying explicit memory as well as hippocampal LTP.

3.2 Implicit Memory in Transgenic Mice CaMKII is also implicated in implicit memory. New molecular technologies can achieve both temporal and spatial control of transgene expression by combining a forebrain-specific promoter with the tetracycline transactivator system. Using such a system with the CaMKIIa promoter, Mayford et al. (1996a) produced transgenic mice in which high levels of CaMKIIa-Asp-286 could be induced in the lateral amygdala and striatum. The mice exhibited severe impairment in contextual cued fear conditioning tasks. Fear conditioning has components of both implicit and explicit learning. The contextual version of the task involves explicit memory and is selectively impaired by hippocampal lesions (Kim and Fanselow, 1992), while both cued and contextual versions of the task require implicit memory and are impaired by lesions of the amygdala. Interestingly, transgenic mice also showed impaired memory consolidation or recall in fear conditioning. Further studies are required to address whether CaMKII autophosphorylation is critical for implicit memory.

4

CaMKII in Developmental Synaptic Plasticity

4.1 Interaction with NMDA Receptor Many synapses in the early postnatal hippocampus have only NMDA receptors postsynaptically and lack functional AMPA receptors, making them silent at resting membrane potential. When these synapses are subjected to LTP-inducing stimuli, excitatory postsynaptic currents (EPSCs) mediated by AMPA receptors appear. CaMKII activation accounts for maturation of synapses from silent to functional even in the adult brain. Upon CaMKII activation/autophosphorylation, CaMKII binds to NMDA receptors within the PSD. NMDA receptors are heteromeric, composed of two NR1 subunits containing glycinebinding sites combined with two NR2 subunits containing glutamate-binding sites. Expression of four NR2 subunit subtypes (NR2A–2D) is developmentally regulated. NR2B and NR2D are highly expressed prenatally and expressed at lower levels postnatally. As their levels decline, NR2A and NR2C levels increase. In the adult mouse brain, NR2A is abundant in most brain regions and NR2D is limited to small numbers of cells in fewer regions. Likewise, NR2B expression is mostly limited to forebrain, whereas NR2C is expressed in the cerebellum (Monyer et al., 1994; Standaert et al., 1996). Importantly, autophosphorylation increases association of CaMKIIa and CaMKIIb with the C-terminus of the NR2B subunit

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(Strack and Colbran 1998). CaMKII also binds the NR1 subunit with relatively lower affinity than to NR2B. Under resting conditions, CaMKIIb binds to F-actin away from the PSD. NMDA receptor stimulation by glutamate during synaptic development triggers CaMKII translocation to the PSD via autophosphorylation. One target molecule for autophosphorylated CaMKII in the PSD is the NR2B subunit (Strack et al., 1997a; 1997b). Association of CaMKII with NR2B also converts the former to a constitutively active form by interaction of NR2B residues 1290–1309 with the CaMKII T-site in an autophosphorylation-independent manner. Maintenance of active CaMKII in the PSD also requires CaMKII binding to two other PSD proteins, densin-180 and a-actinin. The latter promotes CaMKII binding to F-actin, thereby maintaining active CaMKII in the PSD.

4.2 Interaction with MAGUK Proteins Various membrane-associated guanylate kinase (MAGUK)-related proteins, including PSD-95, PSD-93, and synaptic associated protein (SAP)-102, anchor NMDA receptors to the synapse. These proteins exhibit PDZ (PSD-95/Dlg/ZO1) domains, thereby functioning as synaptic scaffold proteins. Among them, PSD-95 interacts with an NR2A/2B N-terminal motif (amino acid residues of T/SXV) through the PDZ domain. Since other signaling molecules including Src and Fyn tyrosine kinases, neuronal nitric oxide synthase (nNOS) and SynGAP bind to PSD-95, these signaling molecules are preferentially activated by Ca2+ mobilization through NMDA receptors through PSD-95 in the PSD. During brain development, the NMDA receptor is localized both synaptically and extrasynaptically. SAP-102 synaptically anchors NR2B as neurons develop prenatally, whereas PSD-95 and PSD-93 levels developmentally increase coincident with NR2A expression in mature synapses during postnatal development. CaMKIIa binds to NR2B with greater affinity than to NR2A, implying that CaMKIIa contributes to synaptic plasticity in immature synapses in early development. The higher Ca2+ and Na+ conductance in NR1/NR2B compared with NR1/NR2A receptors may be required for rapid development of synaptic connectivity and synaptic changes in the prenatal brain.

4.3 Interaction with AMPA Receptor In addition to promoting new synapse formation during development, activity-dependent CaMKII translocation to the PSD is critical for AMPA receptor anchoring and trafficking to synapses, thereby converting silent to functional synapses. AMPA receptors are tetrameric heteromeric complexes of four homologous GluR1–GluR4 (GluRA-GluRD) subunits. Most hippocampal AMPA receptors consist of GluR1/GluR2 and GluR2/GluR3 complexes, whereas most AMPA receptors in cerebellar Purkinje cells are GluR2/GluR3 heteromers (Song and Huganir, 2002). GluR1 binds to other synapse-associated proteins, such as SAP97 (Leonard et al., 1998) and protein 4.1N (Shen et al., 2000). Since the GluR1/SAP97 complex binds to actin filaments through protein 4.1N, actin filaments are a link between the CaMKII/a-actinin complex and a protein 4.1N/SAP97 complex, thereby recruiting AMPA receptor complexes near the PSD formed by NR2B/ CaMKII. NMDA activity or CaMKII-dependent recruitment of AMPA receptors to synapses plays a central role in hippocampal LTP induction. The GluR2 subunit is constitutively delivered to synapses, whereas delivery of GluR1-containing AMPA receptors requires NMDA receptor or CaMKII activation (Shi et al., 1999; Hayashi et al, 2000). Notably, mutation of the PDZ binding motif in the C terminus of GluR1 inhibits activity or CaMKII-dependent delivery of GluR1, implying that PDZ domain-containing proteins like SAP97 and PSD-95 are important for GluR1 delivery to synapses (Hayashi et al., 2000).

4.3.1 Stargazin in GluR1 Delivery Mutations in the gene stargazin cause ataxia and epileptic seizures in the stargazer mouse. These mutations likely underlie CaMKII-dependent delivery of GluR1 (Chen et al., 2000). Stargazin first recruits AMPA receptors from submembranous sites to the plasma membranes, and secondarily stargazin associates with

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PSD proteins to recruit AMPA receptors to synapses (Chen et al., 2000). Phosphorylation of stargazin at Ser-9 by CaMKII and PKC facilitates the second step without effecting the first (Tomita et al., 2005). Conversely, dephosphorylation of that residue by protein phosphatase 1 (PP1) promotes long-term depression in hippocampal synapses. In addition to stargazin, GluR1 is a substrate for CaMKII after delivery by stargazin to the PSD. Ser-831 and Ser-845 in the GluR1 C-terminus are preferentially phosphorylated by CaMKII/PKC and protein kinase A (PKA), respectively (Roche et al., 1996; Barria et al., 1997a; Mammen et al., 1997). Barria et al. (1997) showed that LTP induction increases phosphorylation of GluR1 by CaMKII. Since phosphorylation of GluR1 increases the apparent single-channel conductance of GluR1 (Derkach et al., 1999), GluR1 phosphorylation mediates in part potentiation of AMPA current during LTP induction (Benke et al., 1998). Consistently, in hippocampal CA1 pyramidal neurons, infusion of a constitutively active form of CaMKII via the recording pipette increased the size of EPSCs, as well as the amplitude of responses to iontophoretically applied AMPA (Lledo et al., 1995). Enhancement of AMPA current by CaMKII was greatly diminished by prior induction of LTP. Conversely, following an increase in synaptic strength mediated by CaMKII, tetanic stimulation failed to induce LTP. These findings lead to a simple switch model of LTP in which CaMKII alone is sufficient to induce LTP.

5

Phosphorylation-Dependent Synaptic Plasticity in Mature Synapses

5.1 Molecular Switch of LTP and LTD Phosphorylation and dephosphorylation constitutes a molecular switch between LTP and LTD, respectively, in hippocampal slices (> Figure 9-3). Interestingly, both LTP and LTD events are triggered by NMDA receptor activation at hippocampal CA1 synapses (Malenka and Nicoll, 1999). The amplitude/duration of Ca2+ mobilization through the NMDA receptor and the spatial extent of Ca2+ mobilization at the synapse define the direction of LTP and LTD in glutamatergic synapses. For example, high frequency stimulation greater than 5 Hz at excitatory synapses causes large (greater than 1 mM) and transient Ca2+ elevation in a limited postsynaptic region where CaMKII is preferentially activated in the PSD. On the other hand, prolonged low frequency stimulation of 1 Hz elicits small ( Figure 9-3). Thus protein phosphatase-induced Ser-845 dephosphorylation triggered by low frequency stimulation mediates in part depressed synaptic transmission during hippocampal LTD (Lee et al., 2000). One form of LTD in the CA1 region requires activation of postsynaptic NMDA receptors and results in increases in postsynaptic Ca2+ concentrations. Such an increased Ca2+ concentration, which may be too low to stimulate CaMKII, activates the Ca2+/CaM-dependent protein phosphatase, calcineurin, resulting in dephosphorylation of inhibitor 1. This event leads to increased PP1 activity accounting for a decrease in autonomous CaMKII activity and dephosphorylation of GluR1 Ser-845 at postsynaptic densities. In support of this idea, bath application of FK506, a calcineurin inhibitor, or infusion of phosphorylated inhibitor 1 into postsynaptic cells prevents LTD induction (Mulkey et al., 1994; Morishita et al., 2001). PP1 is targeted to a complex associated with GluR1 that includes SAP97, AKAP150, PKA, PP2B, and inhibitor 1 (Morishita et al., 2001). In addition, during LTD, spinophilin, another PP1 binding protein, mediates PP1 translocation from dendrites to synapses, where spinophilin recruits PP1 to GluR1 complexes. Indeed, spinophilin knockout

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mice do not exhibit LTD (Feng et al., 2000). Thus phosphorylation and dephosphorylation of GluR1 is molecular switch between LTP and LTD in mature synapses. As noted, in contrast with NMDA receptordependent recruitment of GluR1, recruitment of GluR2 subunits is rapid and not regulated by NMDA receptor activity. Phosphorylation of GluR2 Ser-880 promotes AMPA receptor endocytosis during LTD (Xia et al., 2000).

5.4 PKC in Cerebellar LTD PKC activation is critical to induction of cerebellar LTD, in which phosphorylation of Ser-880 by PKC disrupts the interaction of GluR2 with GRIP (glutamate receptor-interacting protein), an AMPA receptor scaffold protein exhibiting a PDZ domain. GluR2 internalization is promoted by PICK1 (protein interacting with C kinase), another GluR2 binding protein (Chung et al., 2002; Matsuda et. al., 2000). Inhibiting the interaction of GluR2 with PICK1 by infusing peptides disrupting GluR2–PDZ interactions into Purkinje cells inhibits cerebellar LTD induction (Xia et al., 2000). Likewise perfusion of those peptides into hippocampal pyramidal neurons inhibits LTD. However, GluR2 Ser-880 phosphorylation in hippocampal LTD is independent of PKC activity (Kim et al., 2001). Taken together, phosphorylation of GluR2 Ser-880 is critical for both hippocampal and cerebellar LTD.

6

LTP Maintenance

6.1 Protein Synthesis-dependent LTP Late-phase LTP is believed to require protein synthesis, since LTP maintenance is associated with sprouting of synapses and/or morphological changes in dendritic spines. Experiments using protein synthesis inhibitors reveal that protein synthesis is necessary to maintain and consolidate LTP and that proteins are synthesized from preexisting rather than newly transcribed mRNAs. Maintenance of increased AMPA responses apparently requires expression of newly synthesized AMPA receptors, a process requiring the cAMP-dependent protein kinase (Nayak et al., 1998). Since the existence of mRNAs encoding CaMKIIa and MAP2 in neuronal dendrites has been demonstrated in hippocampal CA1 regions or in cultured neurons, increased levels of CaMKIIa and MAP2 proteins would be predicted. Increased levels of CaMKII protein have also been confirmed by immunohistochemistry (Ouyang et al., 1997). These changes in levels of total CaMKII protein were observed within 1 h of tetanization. Anisomycin, which inhibits translation, was injected immediately after tetanization and produced a decay in LTP. When injected after a 15 min delay, it had no effect. Taken together, early protein synthesis required for LTP maintenance is mostly complete within 15 min of tetanization. In addition, Mackler et al. (1992) observed a threefold increase in CaMKII mRNA levels between 30 min and 3 h after tetanization.

6.2 Delivery of mRNA to Dendrite Similarly, in the dentate gyrus of freely moving rats, transient CaMKIIa mRNA expression was observed in the soma, and a more persistent increase in CaMKIIa mRNA was observed in the dendritic field without changes in CaMKIIb mRNA following LTP induction (Thomas et al., 1994). Targeting CaMKIIa mRNA to dendrites requires several cis-acting elements in the 30 untranslated region, one of which requires synaptic activity (Mayford et al., 1996; Mori et al., 2000). Protein synthesis-dependent LTP maintenance in the hippocampus requires CaMKII activation through NMDA receptor activation. In addition to targeting of CaMKIIa mRNA to dendrites, CaMKII itself promotes translation of its own mRNA in hippocampal dendrites, where phosphorylation of the cytoplasmic polyadenylation element binding protein (CPEB) regulates protein synthesis. CPEB binds to the 30 untranslated region (UTR) of cytoplasmic mRNAs, and when phosphorylated by CaMKII, initiates mRNA polyadenylation and translation. (Atkins et al., 2004)

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Long-lasting LTP induced by strong stimulation in hippocampal slices leads to prolonged phosphorylation of CPEB by CaMKII, thereby stabilizing synaptic enhancement (Atkins et al., 2005).

7

Morphological Change in Spine

7.1 Spine Formation in LTP In addition to synaptic biochemical changes, morphological changes were also evident in activated dendrites and spines following high frequency stimulation in organotypic cultured hippocampus, while NMDA receptor stimulation leading to LTP in hippocampal slices resulted in the outgrowth of filopodia and spines with a time course of minutes to hours (Hosokawa et al., 1995; Engert and Bonhoeffer, 1999; Maletic-Savatic et al., 1999; Toni et al., 1999). Likewise, LTP induction results in a formation of perforated synapses, followed by creation of multiple spines on the axon terminus in hippocampal slices (Toni et al., 1999). Observed increases in the number of perforated synapses were transient and significant only 30 min after LTP induction. The number of multiple spine boutons gradually increased, reached a maximum at 1 h, and was stable for at least 2 h. Interestingly, both structural remodeling of synapses in LTP were blocked by treatment with KN93, a CaMKII inhibitor. Actin filaments play a pivotal role in synaptic transmission and LTP. For example, bath application of the actin polymerization inhibitor latrunculin A reduced the synaptic response through AMPA receptors (Kim and Lisman, 1999). Infusion of latrunculin A or phalloidin, an actin filament stabilizer, decreased the magnitude of LTP without affecting baseline transmission. Consistent with these results, F-actin depolymerization following treatment with latrunculin A led to a 40% decrease in both the number of synaptic NMDA receptor clusters and the number of AMPA receptor (GluR1)-labeled spines (Allison et al., 1998). Interestingly, CaMKIIb functions as a targeting module that localizes CaMKIIa and CaMKIIb hetero-oligomers to dendritic spines due to specific binding of CaMKIIb to F-actin (Shen et al., 1998). Infusion of constitutively active CaMKII into hippocampal pyramidal neurons markedly enhanced filopodium formation and spine head enlargement (Muller et al., 2003).

7.2 Actin Reorganization Actin reorganization in dendritic spines is critical for spine morphological changes (Fischer et al., 2000; Matus et al., 2000; Smart and Halpain, 2000; Star et al., 2002). Spine shape and motility are regulated by cell surface receptors and ion channels, which activate signaling cascades controlling activity of Rho GTPase and Ca2+ mobilization into spines. Rho GTPase family members, including RhoA, Rac, and Cdc 42, are key elements of actin cytoskeleton formation in spines (Ridley, 2001; Etienne-Manneville and Hall, 2002). In cultured hippocampal and cortical slices, overexpression of constitutively active Rac1 induces formation of irregularly shaped spines, such as lamellipodia-like protrusions (Nakayama et al., 2000; Tashiro et al., 2000; Govek et al., 2004). Overexpression of dominant-negative forms of Rac1 decreases both the number of spines and synapses in cultured hippocampal slices and dissociated hippocampal neurons (Nakayama et al., 2000; Penzes et al., 2003; Zhang et al., 2003). Likewise, dominant-negative Cdc42 inhibits spine morphogenesis (Irie and Yamaguchi, 2002). On the other hand, overexpression of constitutively active Rho A in hippocampal slices promotes spine retraction and elimination, resulting in decreased spine density (Tashiro et al., 2000; Govek et al., 2004). Regarding downstream mechanisms regulating actin dynamics by Rho GTPase, Cdc42 promotes actin nucleation and branching in actin filaments by interaction with the actinrelated 2/3 complex (Arp2/3) (Higgs and Pollard 2001; Irie and Yamaguchi, 2002; Miki et al., 1996).

7.3 PAK Activation Both Rac1 and Cdc42 also activate PAK1, a serine-threonine kinase that phosphorylates and activates LIM kinases (LIMKs) (Yang et al., 1998; Edwards et. al., 1999) (> Figure 9-4). The LIMKs phosphorylate and

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. Figure 9-4 Possible mechanisms underlying morphological and functional modification of spines by CaMKII. CaMKII structurally and enzymatically regulates spine reorganization in developmental and synaptic activitydependent synaptic plasticity. CaMKIIb and CaMKIIa/b hetero-oligomers bind to actin filaments to promote bundling of actin filaments, thereby causing spine enlargement and new spine formation. CaMKII also promotes stability of enlarged spines in activated synapses through F-actin bundling. In addition, phosphorylation of Tiam1 and spinophilin by CaMKII likely mediates activity-dependent spine morphogenesis. Both Rac1 and Cdc42 activate PAK1, a serine-threonine kinase that phosphorylates and activates LIM kinases (LIMKs). LIMKs phosphorylate and inhibit the actin depolymerizating protein cofilin. PAK1 directly phosphorylates myosin light chain (MLC) and in turn activates actomyosin contractility. Taken together, Rho GTPase family members cooperatively regulate actin dynamics in spines, and CaMKII activation by Ca2+ mobilization is critical for spine morphogenesis

inhibit the actin depolymerizating proteins ADF and cofilin. PAK1 directly phosphorylates the myosin light chain (MLC) and in turn activates actomyosin contractility. Since PAK1 also phosphorylates and inactivates myosin light chain kinase, thereby inhibiting MLC phosphorylation, the physiological relevance of MLC phosphorylation by PAK1 remains unclear. PAK3, a related PAK1 family member, is an X-linked mental retardation-related gene highly expressed in brain, particularly in the hippocampus. (Allen et al., 1998). Interestingly, antisense- and siRNA-mediated PAK3 suppression, as well as expression of dominantnegative forms of PAK3, causes marked increases in the proportion of elongated, thin and tortuous dendritic spines and filopodia-like protrusions (Boda et al., 2004). Taken together, Rho GTPase family members cooperatively regulate actin dynamics in spines, and Ca2+ mobilization in synaptic activity likely affects these pathways.

7.4 Ca2+ Mobilization Ca2+ mobilization through NMDA and AMPA receptor channels and other G protein-coupled receptors triggers both developmental morphogenesis and synaptic activity-dependent morphogenesis in spines. The

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magnitude and duration of Ca2+ mobilization have differing effects on spine morphology. Moderate and transient increases in Ca2+ concentration induce spine elongation and formation of new spines (Korkotian and Segel, 1998). By contrast, large and sustained increases in Ca2+ concentration cause spine shortening and collapse (Segal, 1995; Halpain et al., 1998). Several actin binding proteins regulating Ca2+ concentration are localized in spines. Profilin II promotes actin polymerization by binding actin monomers to the barbed ends of growing actin filaments. In cultured hippocampal neurons, NMDA receptor stimulation leads to recruitment of profilin II into the dendritic spine head (Ackermann and Matus, 2003). Gelsolin also caps the barbed ends of actin filaments. Elevated Ca2+ concentrations due to toxic doses of glutamate promote binding of gelsolin to actin, leading to Ca2+-dependent depolymerization of actin filaments in dendritic spines (Furukawa et al., 1997). Spinophilin (also called neurabin II) and neurabin I, both of which promote targeting of PP1 to spines, contribute to bundling of actin filaments in dendritic spines (Nakanishi et al., 1997; Feng et al., 2000). Spinophilin knockout mice have more filopodia and spines early in development and altered glutamatergic transmission (Feng et al., 2000). Phosphorylation of spinophilin by CaMKII and PKA inhibits its binding to actin filaments and keeps AMPA receptors away from the PSD, resulting in decreased GluR1dephosphorylation. These events are associated with stability of the actin cytoskeleton in spines. The opposite effects of neurabin I are evident in spine actin dynamics. Neurabin I overexpression induces formation of dendritic filopodia in immature cultured hippocampal neurons (Oliver et al., 2002).

7.5 Guanine-Nucleotide Exchange Factors A possible downstream target for spinophilin is a guanine-nucleotide exchange factor (GEF) Tiam1, which is implicated in Rac1-mediating actin reorganization. Loss of Tiam1 function mediated by dominantnegative mutants or RNA interference causes marked reduction in dendritic spine density. Tiam1 nucleotide exchange activity toward Rac1 is also stimulated by CaMKII-dependent phosphorylation (Fleming et al., 1999) (> Figure 9-4). Together with Tiam, Kalirin-7, other GEF for Rac1, control activity-dependent dendrite development and synapse maturation. Kalirin-7 interacts with several PDZ domain-containing proteins such as PSD-95, SAP-97, and SAP-102 (Penzes et al., 2001). NMDA receptor activation in pyramidal neurons triggers CaMKII-dependent phosphorylation of kalirin-7 (Thr95), thereby leading to activation of Rac1 and rapid enlargement of existing spines (Xie et al., 2007). Tiam1 mediate the activitydependent increase in spine density during synapse maturation (Tolias et al., 2005), while kalirin-7 accounts for activity-dependent increases in spine size (Xie et al., 2007). Taken together, Both Tiam1 and kalirin-7 are targets for CaMKII activity induced by NMDA receptor activation (> Figure 9-4).

7.6 SynGAP CaMKII itself structurally and enzymatically regulates spine reorganization in a developmental and synaptic activity-dependent manner. CaMKIIb and CaMKIIa/b hetero-oligomers bind to actin filaments to promote bundling of actin filaments, thereby causing spine enlargement and new spine formation (Shen et al., 1998; Jourdain et al., 2003; Pratt et al., 2003). CaMKII also promotes stability of enlarged spines in activated synapses through F-actin bundling (Pratt et al., 2003; Okamoto et al., 2007). In addition to Tiam1 and spinophilin, CaMKII can phosphorylate other signaling proteins associated with activity-dependent spine morphogenesis. SynGAP, a GTPase activating protein, is a particularly attractive candidate as a neuronal target of CaMKII and is a key molecule promoting CaMKII-Ras-ERK pathways following NMDA receptor activation. SynGAP phosphorylation activates Ras and in turn ERK activity. SynGAP null hippocampal cultured neurons form longer spines than do wild-type neurons (Vazquez et al., 2004). Mutation of neurofibromin, another major GTPase activating protein for Ras in neurons, is associated with mental retardation (Costa et al., 2002; Weeber and Sweatt, 2002).

The function of CaM kinase II in synaptic plasticity and spine formation

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7.7 EphB Receptor Because CaMKII functions in the recruitment and/or clustering of AMPA receptors in spines, phosphorylation is implicated in expression of functional AMPA receptors in developmental and activity-dependent synaptic plasticity. During early brain development, NMDA receptors containing NR2B are major components and involved in excitatory glutamatergic transmission. During nervous system development, Eph receptors and ephrin-B ligands play pivotal roles not only in axon guidance but also in establishing appropriate synaptic contact between filopodia and axons. Several EphB receptors are expressed in dendritic spines (Buchert et al., 1999; Grunwald et al., 2001). In immature synapses, soluble ephrin-B1 ligand induces clustering of EphB2 receptors with NMDA receptors and other postsynaptic PDZ domaincontaining scaffold proteins, including GRIP1 and Pick1. Ephrin-B1 ligand-activated EphB receptors stimulate Src kinase activity, thereby increasing NMDA receptor activity. NMDA receptor activation in turn leads to accumulation of NMDA receptor-binding scaffold proteins such as PSD-95. Once NMDA receptors are activated by early synaptic contact between filopodia and axons, autophosphorylated CaMKIIa accumulates and is targeted to the PSD. Prolonged activation of NMDA receptors by repeated synaptic activation promotes accumulation of autophosphorylated CaMKII to stabilize synapses by enlargement of spines, where AMPA receptors are recruited in a synaptic activity-dependent manner. Even in mature synapses in the adult brain, synaptic activity-dependent recruitment of AMPA receptors and morphological spine changes have key roles in learning and memory formation.

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Abnormal Spine Morphogenesis in Mental Retardation

8.1 Genes Associated with Mental Retardation Abnormalities in dendritic spines are associated with deficits in cognitive function and learning. Since it is certain that CaMKII and signaling complexes affect actin dynamics in spine formation, neurodevelopmental disorders characterized by abnormalities spine morphogenesis are likely associated with aberrant signal transduction. Indeed mental retardation seen in conditions such as Fragile X syndrome, William’s syndrome, Rett syndrome, Down’s syndrome, Angelman syndrome, and autism are often accompanied by abnormal shape and density of dendritic spines (Rudelli et al., 1985; Ferrer and Gullotta, 1990; Irwin et al., 2000; Kaufmann and Moser, 2000; Barnes and Milgam, 2002; Fiala et al., 2002; Boda et al., 2004). Several mutant genes associated with mental retardation encode components of Rho family pathways. X-linked mutated genes include Rac1 and the Cdc42 exchange factor aPIX (Kutsche et al., 2000; Govek et al., 2004), the Cdc42 exchange factor faciogenital dysplasia gene 1 (FDG1) (Pasteris et al., 1994; Zheng et al., 1996), the RhoA GTPase activating protein oligophrenin-1 (Billuart et al., 1998), and the Rac1 and Cdc42 downstream effectors, PAK3 (Allen et al., 1998). Other forms of mental retardation such as Angelman syndrome, Rett syndrome, and ATR-X (X-linked a-thalassemia/mental retardation) syndrome are not associated with these types of mutations. Thus mechanisms underlying dendritic spine abnormalities seen in a subset of mental retardation conditions remain unclear.

8.2 CaMKII in Angelman and ATR-X Syndromes Interestingly, a mouse model for Angelman syndrome exhibiting a Ube3a maternal null mutation shows reduced CaMKII activity, autophosphorylation capability, and total CaMKII associated with the PSD (Weeber et al., 2003). Importantly, neurological deficits observed in the Angelman syndrome mouse are rescued by expression of active CaMKIIa (von Woerden et al., 2007). Rett syndrome is caused by mutations in the methyl-CpG-binding protein 2 (MECP2) gene (Amir et al., 1999), encoding a long-range transcriptional repressor that binds to methylated DNA. Phosphorylation of MeCP2 at Ser-421 by CaMKII stimulates spine morphogenesis and activity-dependent induction of BDNF (Zhou et al., 2006). ATR-X

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syndrome is also induced by abnormal chromatin remodeling. ATR-X model mice generated by a splicing mutation show abnormal CaMKII activity in the prefrontal cortex, where abnormal phosphorylation of Tiam1 and PAK3 are also observed (Shioda et. al., 2007). Taken together, although causative mechanisms underlying abnormal CaMKII activity in Angelman syndrome and ATR-X syndrome model mice are unclear, dysregulation of CaMKII activity may trigger the abnormalities of spine morphology in the mental retardation.

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Conclusion

CaMKII is critical molecule not only for developmental synaptic maturation but also synaptic plasticity in the mature brain. In the later case, CaMKII activity is required for memory and learning functions in mammalian brains. Synaptic activity-dependent phosphorylation and trafficking of the GluR1 subunit into the PSD through CaMKII activation account for the hippocampal LTP as a molecular basis of memory. Moreover, CaMKII activation in part mediates rapid morphological changes in dendritic spines at excitatory synapses during LTP induction. In addition to the critical role played by CaMKII in synaptic plasticity in various brain regions, various psychotic disorders including mental retardation, schizophrenia, and depression alter CaMKII activity in specific brain regions correlated with changes in spine morphology. The extensive studies are required to define neuronal mechanism eliciting dysregulation of CaMKII in the specific brain regions during process of mental retardation.

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Penzes P, Johnson RC, Sattler R, Zhang X, Huganir RL, et al. 2001. The neuronal Rho-GEF Kalirin-7 interacts aith PDZ domain-containing proteins and regulates dendritic morphogenesis. Neuron 29: 229-242. Pettit DL, Perlman S, Malinow R. 1994. Potentiated transmission and prevention of further LTP by increased CaMKII activity in postsynaptic hippocampal slice neurons. Science 266: 1881-1885. Pratt KG, Watt AJ, Griffith LC, Nelson SB, Turrigiano GG. 2003. Activity-dependent remodeling of presynaptic inputs by postsynaptic expression of activated CaMKII. Neuron 39: 269-281. Ridley AJ. 2001. Rho family proteins: Coordinating cell responses. Trends Cell Biol 11: 471-477. Roche KW, O’Brien RJ, Mammen AL, Bernhardt J, Huganir RL. 1996. Characterization of multiple phosphorylation sites on the AMPA recetor GluR1 subunit. Neuron 16: 1179-1188. Rotenberg A, Mayford M, Hawkins R, Kandel ER, Muller RU. 1996. Mice expressing activated CaMKII lack low frequency LTP and do not form stable place cells in the CA1 regions of the hippocampus. Cell 87: 1351-1361. Rudelli RD, Brown WT, Wisniewski K, Jenkins EC, LaureKamionowska M, et al. 1985. Adult fragile X syndrome. Clinico-neuropathologic findings. Acta Neuropathol (Berl) 67: 289-295. Schulman H, Hanson PI. 1993. Multifunctional Ca2+/ calmodulin-dependent protein kinase. Neurochem Res 18: 65-77. Schworer CM, Colbran RJ, Keefer JR, Soderling TR. 1988. Ca2+/calmodulin-dependent protein kinase II: Identification of a regulatory autophosphorylation site adjacent to the inhibitory and calmodulin-binding domains. J Biol Chem 263: 13486-13489. Schworer CM, Rothblum LI, Thekkumkara TJ, Singer HA. 1993. Identification of novel isoforms of the d subunit of Ca2+/calmodulin-dependent protein kinase II. Differential expression in rat brain and aorta. J Biol Chem 268: 14443-14449. Segal M. 1995. Morphological alterations in dendritic spines of rat hippocampal neurons exposed to N-methyl-Daspartate. Neurosci Lett 193: 73-76. Shen L, Liang F, Walensky LD, Huganir RL. 2000. Regulation of AMPA receptor GluR1 subunit surface expression by a 4.1N-linked actin cytoskeletal association. J Neurosci 20: 7932-7940. Shen K, Teruel MN, Subramanian K, Meyer T. 1998. CaMKIIb functions as an F-actin targeting module that localizes CaMKIIab heterooligomers to dendritic spines. Neuron 21: 593-606. Shi SH, Hayashi Y, Petralia RS, Zaman SH, Wenthold RJ, et al. 1999. Rapid spine delivery and redistribution of AMPA

receptors after synaptic NMDA receptor activation. Science 284: 1811-1816. Shioda N, Tokoro T, Kitajima I, Beppu H, Fukunaga K. 2007. Activation of CaM kinase II is implicated in generation of abnormal dendritic spines via Tiam1/PAK signaling in ATRX mutant mouse brain. Abstr Soc Neurosci Silva AJ, Paylor R, Wehner JM, Tonegawa S. 1992b. Impaired spatial learning in alpha-calcium-calmodulin kinase II mutant mice. Science 257: 206-211. Silva AJ, Stevens CF, Tonegawa S, Wang Y. 1992a. Deficient hippocampal long-term potentiation in alpha-calciumcalmodulin kinase II mutant mice. Science 257: 201-206. Smart FM, Halpain S. 2000. Regulation of dendritic spine stability. Hippocampus 10: 542-554. Soderling TR, Chang B, Brickey D. 2001. Cellular signaling through multifunctional Ca2+/calmodulin-dependent protein kinase II. J Biol Chem 276: 3719-3722. Song I, Huganir RL. 2002. Regulation of AMPA receptors during synaptic plasticity. Trends Neruosci 25: 578-588. Srinivasan M, Edman CF, Schulman H. 1994. Alternative splicing introduces a nuclear localization signal that targets multifunctional CaM kinase to the nucleus. J Cell Biol 126: 839-852. Standaert DG, Landwehrmeyer GB, Kerner JA, Penney JB Jr, Young AB. 1996. Expression of NMDAR2D glutamate receptor subunit mRNA in neurochemically identified interneurons in the rat neosriaum, neocortex and hippocampus. Brain Res Mol Brain Res 42: 89-102. Star EN, Kwiatkowski DJ, Murthy VN. 2002. Rapid turnover of actin in dendritic spines and its regulation by activity. Nat Neurosci 5: 239-246. Strack S, Barban MA, Wadzinski BE, Colbran RJ. 1997a. Differential inactivation of postsynaptic density-associated and soluble Ca2+/calmodulin-dependent protein kinase II by protein phosphatase 1 and 2A. J Neurochem 68: 2119-2128. Strack S, Colbran RJ. 1998. Auotphosphorylation-dependent targeting of calcium/calmodulin-dependent protein kinase II by the NR2B subunit of N-methyl-D aspartate receptor J Biol Chem 273: 20689-20692. Sweatt JD. 2004. Mitogen-activated protein kinases in synaptic plasticity and memory. Curr Opin Neurobiol 14: 311-317. Takeuchi Y, Fukunaga K, Miyamoto E. 2002. Activation of nuclear Ca2+/Calmoclulin-dependent protein kinase II and brain-derived neutrophic factor gene expression by stimulation of dopamine D2 receptor in transfected NG 108-15 cells. J Neurochem 82: 316-328. Takeuchi Y, Yamamoto H, Fukunaga K, Miyakawa T, Miyamoto E. 2000. Identification of the isoforms of Ca2+/calmodulin-dependent protein kinase II in rat astrocytes and their subcellular localization. J Neurochem 74: 2557-2567.

The function of CaM kinase II in synaptic plasticity and spine formation Takeuchi Y, Yamamoto H, Matsumoto K, Kimura T, Katsuragi S, et al. 1999. Nuclear localization of the d subunit of Ca2+/calmodulin-dependent protein kinase II in rat cerebellar granule cells. J Neurochem 72: 815-825. Tashiro A, Minden A, Yuste R. 2000. Regulation of dendritic spine morphology by the rho family of small GTPases: Antagonistic roles of Rac and Rho. Cereb Cortex 10: 927-938. Thiel G, Czernik AJ, Gorelick F, Nairn AC, Greengard P. 1988. Ca2+/calmodulin-dependent protein kinase II: Identification of threonine-286 as the autophosphorylation site in the a subunit associated with the generation of Ca2+-independent activity. Proc Natl Acad Sci USA 85: 6337-6341. Thomas KL, Laroche S, Errington ML, Bliss TVP, Hunt SP. 1994. Spatial and temporal changes in signal transduction pathways during LTP. Neuron 13: 737-745. Tobimatsu T, Fujisawa H. 1989. Tissue-specific expression of four types of rat calmodulin-dependent protein kinase II mRNAs. J Biol Chem 264: 17907-17912. Tobimatsu T, Kameshita I, Fujisawa H. 1988. Molecular cloning of the cDNA encoding the third polypeptide (g) of brain calmodulin-dependent protein kinase II. J Biol Chem 263: 16082-16086. Tolias KF, Bikoff JB, Burette A, Paradis S, Harrar D, et al. 2005. The Rac1-GEF Tiam1 couples the NMDA receptor to the activity-dependent development of dendritic arbors and spines. Neuron 45: 525-538. Tomita S, Stein V, Stocker TJ, Nicoll RA, Bred DS. 2005. Bidirectional synaptic plasticity regulated by phosphorylation of stargazing-like TARPs. Neuron 45: 269-277. Toni N, Buchs PA, Nikkonenko I, Bron CR, Muller D. 1999. LTP promotes formation of multiple spine synapses between a single axon terminal and a dendrite. Nature 402: 421-425. Vazquez LE, Chen HJ, Sokolova I, Knuesel I, Kennedy MB. 2004. SynGAP regulates spine formation. J Neurosci 24: 8862-8872.

9

von Woerden GM, Harris KD, Hojjati MR, Gustin RM, Qiu S, et al. 2007. Rescue of neurological deficits in a mouse model for Angelman syndrome by reduction of a CaMKII inhibitory phosphorylation. Nat Neurosci 10: 280-282. Weeber EJ, Jiang YH, Elgersma Y, Varga AW, Carrasquillo Y, et al. 2003. Derangements of hippocampal calcium/calmodulindependent protein kinase II in a mouse model for Angelman mental retardation syndrome. J Neurosci 23: 2634-2644. Weeber EJ, Sweatt JD. 2002. Molecular neurobiology of human cognition. Neuron 33: 845-848. Xia J, Chung HJ, Wihler W, Huganir RL, Linden DJ. 2000. Cerebellar long-term depression requires PKCregulated interactions between GluR2/3 and PDZ domain-containing proteins. Neuron 28: 499-510. Xie Z, Srivastava DP, Photowala H, Kai L, Cahill ME, et al. 2007. Kalirin-7 controls activity-dependent structural and functional plasticity of dendritic spines. Neuron 56: 640-656. Yang N, Higuchi O, Ohashi K, Nagata K, Wada A, et al. 1998. Cofilin phosphorylation by LIM-kinase 1 and its role in Rac-mediated actin reorganization. Nature 393: 809-812. Zhang H, Webb DJ, Asmussen H, Horwitz AF. 2003. Synapse formation is regulated by the signaling adaptor GIT1. J Cell Biol 161: 131-142. Zheng Y, Fischer DJ, Santos MF, Tigyi G, Pasteris NG, et al. 1996. The faciogenital dysplasia gene product FGD1 functions as a Cdc42Hs-specific guanine-nucleotide exchange factor. J Biol Chem 271: 33169-33172. Zhou Z, Hong EJ, Cohen S, Zhao WN, Ho HY, et al. 2006. Brain specific phosphorylation of MeCP2 regulates activity-dependent Bdnf transcription, dendritic growth, and spine maturation. Neuron 52: 255-269.

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T. Ohshima . K. Mikoshiba

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186

2

Cdk5 Is a Unique Member of the Cdk Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186

3 3.1 3.2 3.3 3.4

Regulation of CDK5 Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186 Heterodimers of Cdk5/p35 and Cdk5/p39 Exhibit Cdk5 Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 186 Phosphorylation of Cdk5 May Regulate In Vivo Cdk5 Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 Expression and Subcellular Localization of p35 and p39 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 Degradation of p35 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 190

4 4.1 4.2 4.3 4.4

Cdk5 Function in Brain Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 190 Role of Cdk5 in Neuronal Migration and Positioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 190 Role of Cdk5 in Neurite Outgrowth, Dendrite Growth, and Spine Development . . . . . . . . . . . . . . . 191 Role of Cdk5 in Sema3A Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191 Role of Cdk5 in Neuronal Survival . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191

5 5.1 5.2 5.3

Cdk5 Function in the Adult Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192 Presynaptic Roles of Cdk5 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192 Postsynaptic Roles of Cdk5 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192 Regulation of Dendritic Spine Remodeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192

6 Role of Cdk5 in Higher Cognitive Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 6.1 Learning Memory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 6.2 Regulation of Dopaminergic Signaling and Drug Addiction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 7 Cdk5 Function in the Peripheral Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 7.1 Role of Cdk5 in Pain Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 8 8.1 8.2 8.3

Cdk5 Function Outside the CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194 Myogenesis and NMJ Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194 Secretory Exocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194 Insulin Secretion in Pancreatic b-Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194

9 9.1 9.2 9.3 9.4

Deregulation of Cdk5 and Neurodegenerative Disorder . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194 Production of p25 Changes Stability and Localization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194 p25 and AD . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195 Cdk5 in Ischemic Brain Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195 Deregulation of Cdk5 and Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195

10

Cdk5 as a Drug Target . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196

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2009 Springer ScienceþBusiness Media, LLC.

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Abstract: Cyclin-dependent kinase 5 (Cdk5) is a neuron-specific serine (Ser)/threonine (Thr) kinase. Accumulating evidence indicates that Cdk5 plays important roles not only in CNS development but also in higher brain function through the phosphorylation of neuronal substrates. In addition, the upregulation of Cdk5 activity is implicated in neurodegenerative disorders such as Alzheimer’s disease. Recent studies also indicate other roles of Cdk5 besides those in the CNS, for example, insulin secretion from b-cells in the pancreas. The possible link between Cdk5 and human disorders indicates that Cdk5 could possibly be used as a drug target. List of Abbreviations: AD, Alzheimer’s disease; APP, amyloid precursor protein; CNS, central nervous system; Cdk5-DN, Cdk5 dominant-negative; Cdk5, Cyclin-dependent kinase 5; GEF, GTP exchange factor; KO, knockout; LTP, long-term potentiation; L-VDCC, L-type voltage-dependent Ca2+ channel; MT, microtubule; NRG, neuregulin; NMJ, neuromuscular junction; RTK, receptor-type tyrosine kinase; TH, tyrosine hydroxylase

1

Introduction

After the discovery of cyclin-dependent kinase 5 (Cdk5) as a neuron-specific kinase in the early 1990s, substantial progress has been made in the identification of its function along with a growing list of substrate proteins (> Table 10‐1). Phenotypic analyses of mice lacking Cdk5 or the Cdk5-activating subunit p35 provide remarkable opportunities for understanding the in vivo function of Cdk5. Cdk5 activity in cells is tightly regulated under physiological conditions. However, the cleavage of p35 to p25 by calpain under pathological conditions changes the localization of active Cdk5, prolongs its stability, and results in the upregulation of its activity. This deregulation of Cdk5 may occur in neurodegenerative disorders such as Alzheimer’s disease (AD) and Parkinson’s disease. In this chapter, we will describe the features of this protein kinase and its function in the developing and mature central nervous system (CNS). Finally, we will discuss the mechanism of Cdk5 deregulation, its role in neurodegenerative disorders, and a prospective view of Cdk5 as a drug target.

2

Cdk5 Is a Unique Member of the Cdk Family

Cdk5, a proline-directed serine (Ser)/threonine (Thr) kinase, had been identified as a member of the CDK family because of its close sequence homology to human CDC2 (Hellmich et al., 1992; Lew et al., 1992; Meyerson et al., 1992). Other Cdks are activated by cyclin in dividing cells and are involved in cell cycle regulation. Cdk5 bound to cyclin D, however, exhibits no kinase activity. Cdk5 is expressed in all tissues (Tsai et al., 1993), although the highest expression and associated kinase activity are detected in the nervous system (Tsai et al., 1993). Monomeric Cdk5 demonstrates no kinase activity and requires association with an activating partner. Two activating subunits of Cdk5—p35 and p39—were identified (Lew et al., 1994; Tsai et al., 1994; Tang et al., 1995). The Cdk5/p35 and Cdk5/p39 heterodimers exhibit in vitro kinase activity (Lew et al., 1994; Tang et al., 1995). Cdk5 and p35/p39 seem to be evolutionally conserved, and orthologues of Cdk5 and p35/p39 have been identified in other vertebrates, invertebrates, and Saccharomyces cerevisiae (Dhavan and Tsai, 2001). Cdk5 phosphorylates the consensus sequence (S/T)PX(K/H/R), where S and T are the phosphorylatable Ser and Thr, respectively, X is any amino acid, and P is the proline in the +1 position (Beaudette et al., 1993; Songyang et al., 1996). Cdk1 and Cdk2 have substrate specificities identical to Cdk5 (Moreno and Nurse, 1990).

3

Regulation of CDK5 Activity

3.1 Heterodimers of Cdk5/p35 and Cdk5/p39 Exhibit Cdk5 Activity Cdk5 binds to one of its activating subunits—p35 and p39—for complete activation. No additional Cdk5 phosphorylation is required for its activation, although other Cdks require phosphorylation of the

Cyclin-dependent kinase 5

10

. Table 10-1 Cdk5 substrates

Biochemical Developmental

Function Protein name Activation of kinase p35 (Cdk5r1) activity Neuronal Doublecortin migration Ndel1

p27

FAK

b-catenin

Filamin1 Dab1 Neurite extension

MAP1b

Tau PAK1

NFH/NFM TrkB

Brain function

Neuronal survival

MEK1

Axon guidance

JNK3 CRMP2

Neurotransmission

Synapsin I

Function of phosphorylation Promoting proteasomal degradation Reducing the binding of Ndel1 to MTs, facilitating neuronal migration Altering Ndel distribution, likely facilitating neuronal migration Maintaining p27 protein level and facilitating neuronal migration via cofilin-mediated actine polymerization Organizing MT network and facilitating nucleokinesis during neuronal migration Regulating cell adhesion and reducing its binding to presenilin-1 Regulating actin dynamics Functional significance is unknown Enhancing MTs binding and neurite extension Reducing the binding of Tau to MTs Regulating actin and microtubule dynamics NF stability and its interaction with MTs Regulating BDNF-induced dendritic growth Inhibiting MEK1 activity Inhibiting JNK3 activity Reducing the binding of CRMPs to MTs, facilitating Sema3A-induced growth cone collapse of DRG neurons Regulating synaptic transmission

References Patrick et al. (1998) Tanaka et al. (2004) Niethammer et al. (2000); Sasaki et al. (2000) Kawauchi et al. (2006)

Xie et al. (2003)

Kwon et al. (2000); Kesavapany et al. (2001) Fox et al. (1998) Keshvara et al. (2002) Paglini et al. (1998); Pigino et al. (1997) Patrick et al. (1999) Nikolic et al. (1998); Rashid et al. (2001) Sun et al. (1996) Cheung et al. (2007) Sharma et al. (2002) Li et al. (2002) Cole et al. (2004); Uchida et al. (2005)

Matsubara et al. (1996)

187

188

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Cyclin-dependent kinase 5

. Table 10-1 (continued) Function

Protein name Munc18

Amphiphysin I

Dynamin I

NR2A P/Q-type calcium channel TH

Synaptojanin Ephexin 1

WAVE1

PSD-95

Signal transduction

a-chimerin RasGRF2 DARPP-32 PP1 inhibitor I-1 Pg (retinal cGMP phosphodiesterase) pRb

p53 STAT3 Peripheral nervous

Pain

TRPV1

Function of phosphorylation Disrupting Munc18Syntaxin1A interaction and regulating neurosecretion Regulating synaptic vesicle endocytosis

References Shuang et al. (1998); Fletcher et al. (1999) Floyd et al. (2001); Tan et al. (2003); Tomizawa et al. (2003) Regulating synaptic vesicle Tan et al. (2003); endocytosis Tomizawa et al. (2003) Regulating synaptic Li et al. (2001) transmission and plasticity Regulating Tomizawa et al. neurotransmitter release (2002) Kansy et al. (2004); Moy and Tsai (2004) Lee et al. (2004) Fu et al. (2007) Regulating EphA4mediated dendritic spine retraction Regulating actin Kim et al. (2006) polymerization and spine morphology Morabito et al. Suppressing PSD-95 (2004) multimerization and reducing PSD-95 dependent clustering of NR1 Possibly regulating Rac1 Qi et al. (2004) activity Suppressing Rac1 activity Kesavapany et al. (2004) Regulating dopamine Bibb et al. (1999) signaling Activating I-1to mediate Huang and PP1 activity Paudel (2000) Regulating retinal Hayashi et al. phototransduction (2000); Matsuura et al. (2000) Possibly regulating Lee et al. (1997) neuronal differentiation and apoptotic death Mediating p53 Zhang et al. transcriptional activity (2002) Mediating STAT3 Fu et al. (2004) transcriptional activity Mediating pain signaling Pareek et al. (2007)

Function

Protein name

Function of phosphorylation

Cyclin-dependent kinase 5

References

10

. Table 10-1 (continued)

Outside of nervous system

Neurodegenerative disorders

Function NMJ

Protein name ErbB

Insulin secretion

L-VDCC

Endocrine secretion Alzheimer’s disease Parkinson’s disease

Trio b-APP MEF2

Prx2 Parkin

Function of phosphorylation Mediating neuregulin signaling at NMJ Inhibiting insulin secretion in high glucose

References Fu et al. (2001) Wei et al. (2005a) Xin et al. (2004)

Mediating APP localization and function Inhibiting MEF2 transcriptional activity and mediating neuronal apoptosis Reducing its peroxidase activity Regulating its ubiquitinligase activity and aggregation

Iijima et al. (2000) Gong et al. (2003)

Qu et al. (2007) Avraham et al. (2007)

activation loop at residue Thr160 for maximal activation. Crystal structure analysis of Cdk5/p25 revealed that this interaction with the activating subunit is sufficient to stretch the activation loop of unphosphorylated Cdk5 into active conformation, thereby rendering it indistinguishable from the phosphorylated Cdk2–cyclin A complex (Tarricone et al., 2001). Cdk5 activity is therefore correlated with the expression levels of p35 and p39. Several p35-binding proteins such as C42 and protein kinase CK2 have been reported, and it has been shown that they inhibit Cdk5 activation by binding to p35 (Ching et al., 2002; Lim et al., 2004).

3.2 Phosphorylation of Cdk5 May Regulate In Vivo Cdk5 Activity It has been reported that Tyr15 of Cdk5 is phosphorylated by c-Abl (Zukerberg et al., 2000). Tyr15 phosphorylation has been shown to be induced by Sema3A during growth cone collapse by the Src family of tyrosine kinases such as Fyn (Sasaki et al., 2002). Recently, BDNF was also shown to induce phosphorylation of Tyr15 in Cdk5 via TrkB, a tyrosine kinase of the BDNF receptor (Cheung et al., 2007). In both cases, Cdk5 activity increased with the phosphorylation of its Tyr15 (Sasaki et al., 2002; Cheung et al., 2007). Neurotransmitters such as glutamate also enhance Cdk5 activity, although the associated Tyr15 phosphorylation is not clearly detected (Wei et al., 2005b). The mechanism by which Cdk5 activity is enhanced by Tyr15 phosphorylation remains to be elucidated.

3.3 Expression and Subcellular Localization of p35 and p39 p35 and p39 are strongly expressed in the postmitotic neurons of the nervous system (Zheng et al., 1998). During brain development, the expression of p35 and p39 overlaps throughout the CNS, except in the cerebral cortex in the early stage where only p35 is expressed until around E16 (Ohshima et al., 2001). High Cdk5 activity during neuronal differentiation and brain development reflects the high-level expression of p35 and p39. The levels of Cdk5 and p35 are transcriptionally regulated. The extracellular matrix glycoprotein laminin increases the p35 transcription levels in cultured cerebellar macroneurons and SH-SY5Y

189

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neuroblastoma cells (Paglini et al., 1998; Li et al., 2000). Nerve growth factor induces p35 expression in PC12 cells via Egr1, a member of the zinc-finger transcription factor family (Harada et al., 2001). Brainderived neurotrophic factor also induces Cdk5 kinase activity (Tokuoka et al., 2000). Chronic cocaine administration to rats has been shown to upregulate Cdk5 and p35 expression via DfosB (Bibb et al., 2001). Cell fraction studies indicate that p35 and p39 localize to overlapping but distinct subcellular compartments in the growth cone and synapses (Humber et al., 2000). Immunocytochemical studies show that p35 is localized at cell periphery such as lamellipodia and filopodia, and p35 and p39 are enriched in membrane fractions (Nikolic et al., 1996). An amino-terminal myristoylation signal motif in p35 and p39 targets them to cellular membranes (Patrick et al., 1999).

3.4 Degradation of p35 The p35 protein is unstable in the neurons, and its half-life was estimated to be 20–30 min by pulse-chase experiments (Patrick et al., 1998). The degradation of p35 is mediated by the ubiquitin/proteasome system, and its phosphorylation by Cdk5 is a signal for ubiquitination/degradation (Patrick et al., 1998; Saito et al., 1998). These mechanisms may function as negative feedback regulation of Cdk5 activity and contribute to its stringent regulation in the neurons. p35 is also cleaved by calpain and converted into the stable p25 protein, as described later.

4

Cdk5 Function in Brain Development

4.1 Role of Cdk5 in Neuronal Migration and Positioning Gene-targeting experiments clearly demonstrate that Cdk5 activity is essential for establishing the architecture of the CNS. Defects in the migration of the cortical neurons of Cdk5-knockout (KO) mice result in the disruption of the laminar structures in the cerebral cortex, olfactory bulb, hippocampus, and cerebellum (Ohshima et al., 1996; Gilmore et al., 1998). p35-KO mice display a milder phenotype than Cdk5-KO mice because of the redundancy of p39 (Chae et al., 1997; Ohshima et al., 2001). p39-KO mice display no phenotype, but p35/p39 double-null mice display a phenotype identical to that of Cdk5-KO mice (Ko et al., 2001), confirming the redundancy of these subunits. Neuronal migration defects in Cdk5-null mice are observed in a subset of neurons, indicating the presence of Cdk5-dependent and Cdk5-independent neuronal migration (Gilmore et al., 1998; Ohshima et al., 2002). For example, in the cerebral cortex, the migration of layer II–V pyramidal neurons is considered Cdk5 dependent, whereas that of either the deepest cortical plate neurons or GABAergic neurons from the ganglionic eminences is Cdk5 independent (Gilmore and Herrup, 2001). Additionally, Cdk5 deficiency in migrating neurons may be compensated by other kinases such as Cdc2. How does Cdk5 regulate neuronal migration and localization? Our recent analysis of migratory abnormality in Cdk5-deficient cortical neurons indicates that Cdk5 is specifically required for multipolarto-bipolar transition, which is a characteristic morphological change in neurons migrating radially in the developing cerebral cortex (Ohshima et al., 2007b). This morphological change in the neurons requires dynamic changes in the cytoskeletal proteins actin and microtubules. Cdk5 modulates actin cytoskeleton dynamics via the phosphorylation of Pak1 (Nikolic et al., 1998; Rashid et al., 2001) and filamin 1 (Fox et al., 1998). It modulates microtubule dynamics via the phosphorylation of microtubule-associated proteins including tau (Kobayashi et al., 1993), MAP1b (Paglini et al., 1998), doublecortin (Tanaka et al., 2004), and Ndel1 (Niethammer et al., 2000; Sasaki et al., 2000). Ndel1 forms protein complex with Lis1 and cytoplasmic dynein (Niethammer et al., 2000; Sasaki et al., 2000). A defect in one of these substrates—filamin 1 in humans—causes periventricular heterotopia (Fox et al., 1998), and defects in two other substrates— LIS1 and doublecortin in humans—cause lissencephaly type 1 (Reiner et al., 1993; des Portes et al., 1998; Gleeson et al., 1998). These findings indicate that Cdk5 regulates cytoskeletal dynamics that determines the

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speed of migration, extension of the leading processes, and cell-soma propulsion in migrating neurons and also regulates proper multipolar-to-bipolar morphological transition. Cdk5 may also mediate cellular adhesion in neuronal–glial interactions through the phosphorylation of b-catenin, thereby regulating its interaction with N-cadherin (Kwon et al., 2000); further, this Cdk5-mediated adhesion may be important for neuronal migration along radial glial fibers. Indeed, less association between migrating neurons and radial glial fibers was reported in p35-deficient mice (Gupt et al., 2003). Although Cdk5 phosphorylates Dab1 at Ser491 in vivo (Keshvara et al., 2002) and at multiple sites in vitro (Ohshima et al., 2007a), the functional significance of Cdk5-mediated Dab1 phosphorylation in Reelin signaling remains to be elucidated (Ohshima and Mikoshiba, 2002). The possible substrates of Cdk5 that are related to neuronal migration and localization are summarized in > Table 10‐1.

4.2 Role of Cdk5 in Neurite Outgrowth, Dendrite Growth, and Spine Development In cultured neurons, the reduction of Cdk5 activity due to the expression of Cdk5 dominant-negative (Cdk5-DN) or the antisense oligonucleotides of Cdk5, p35, and p39 inhibits neurite outgrowth (Nikolic et al., 1996; Xiong et al., 1997). In cerebellar macroneurons, Cdk5 suppression reduced axonal elongation (Pigino et al., 1997; Paglini et al., 1998). These results indicate that Cdk5 activity positively regulates neurite extension. The involvement of Cdk5 in BDNF-stimulated dendritic growth in cultured hippocampal neurons has recently been reported (Cheung et al., 2007). BDNF, a neurotrophin, binds to the receptortype tyrosine kinase (RTK) TrkB and promotes an increase in the number of dendrites among cultured hippocampal neurons; further, this effect does not occur in Cdk5-deficient neurons (Cheung et al., 2007), indicating that Cdk5 is required for BDNF-induced dendritic growth. Cheung et al. (2007) found that Cdk5 phosphorylates TrkB at Ser478 and that the introduction of an S478A TrkB mutant into hippocampal neurons prevents BDNF-induced dendritic growth. We observed abnormal dendritic development in cortex-specific Cdk5-KO mice (Ohshima et al., 2007b). We consider that the morphological changes accompanying radial neuronal migration are disturbed in these mice, and this results in abnormal dendrites in the postnatal cortex (Ohshima et al., 2007b). Therefore, the role of Cdk5 in in vivo dendrite and spine development remains to be elucidated.

4.3 Role of Cdk5 in Sema3A Signaling Cdk5 is also involved in intracellular Sema3A signaling (Sasaki et al., 2002). Sema3A is a repulsive guidance molecule. It causes not only growth cone collapse but also various cellular responses including axonal transport, endocytosis, and spine maturation. Growth cone collapse induced by Sema3A is attenuated in Cdk5-deficient cultured DRG neurons (Sasaki et al., 2002). CRMP2, a mediator of intracellular Sema3A signaling, is phosphorylated by Cdk5 at Ser522, and this phosphorylation is required for the further phosphorylation of CRMP2 at 509/514/518 Ser/Thr sites by GSK3b (Cole et al., 2004; Uchida et al., 2005). A collapse assay with cultured DRG neurons demonstrates that these phosphorylation events are required during Sema3A-induced growth cone collapse (Uchida et al., 2005). These results indicate that Cdk5 participates in Sema3A signaling by phosphorylating CRMP family members.

4.4 Role of Cdk5 in Neuronal Survival Cdk5 promotes neuronal survival by phosphorylating and inactivating JNK3, one of the key players in the apoptotic pathway (Li et al., 2002), and by activating the neuregulin (NRG)/PI3K/Akt survival pathway (Li et al., 2003a). Cdk5 is involved in cross-talk interactions with several intracellular signaling pathways, including the MAPK pathway (Sharma et al., 2002). Cdk5 phosphorylates MEK1 and inhibits Erk1/2 activation (Sharma et al., 2002). Suppression of Cdk5 by siRNA induced an increase in Erk1/2 and neuronal

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apoptosis in cultured cortical neurons, and these effects were eliminated by the MEK1 inhibitor PD98095, indicating that proper Cdk5 activity is required for neuronal survival (Zheng et al., 2007). Cdk5-deficient cultured neurons exhibit increased sensitivity to apoptotic stimuli (Li et al., 2002). An increase in the number of apoptotic cells in the cerebral cortex of Cdk5-KO mice was reported (Li et al., 2002), although increased neuronal death has not been quantitatively evaluated thus far.

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Cdk5 Function in the Adult Brain

Accumulating evidence supports the function of Cdk5 in neurotransmission and synaptic plasticity (see review by Cheung et al., 2006). A myriad of pre- and postsynaptic proteins have been identified as Cdk5 substrates (> Table 10‐1). Recent studies also suggest the pivotal role of Cdk5 in the regulation of higher cognitive functions.

5.1 Presynaptic Roles of Cdk5 By phosphorylating presynaptic proteins including synapsin I (Matsubara et al., 1996), Munc18 (Shuang et al., 1998; Fletcher et al., 1999), and those involved in the P/Q subtype voltage-dependent calcium channel (Tomizawa et al., 2002), Cdk5 is associated with the regulation of synaptic vesicle exocytosis. Cdk5 also phosphorylates dynamin I and amphiphysin I, which are proteins essential for clathrin-mediated endocytosis (Tan et al., 2003; Tomizawa et al., 2003). Cdk5 phosphorylates synaptojanin 1, a polyphosphoinositide phosphatase (Lee et al., 2004). These findings indicate that Cdk5 functions as a negative regulator of neurotransmitter release. Cdk5 has been implicated in the inhibition of dopamine release in the striatum (Chergui et al., 2004). In Cdk5-KO mice, the frequency of the miniature endplate potential is increased (Fu et al., 2005). These experimental results support the view of the inhibitory role of Cdk5 in in vivo neurotransmitter release.

5.2 Postsynaptic Roles of Cdk5 Cdk5 affects the expression and clustering of neurotransmitter receptors at the synapses. This finding was reported with regard to the neuromuscular junction (NMJ) (Fu et al., 2001) as discussed later, and the same findings were observed with regard to the CNS synapses. NRG-induced activation of the NRG receptor ErbB4 results in the Cdk5-dependent upregulation of GABAA receptor transcription in cultured granule cells (Xie et al., 2004). The in vivo significance of these findings by using a Cdk5-deficient mouse model remains to be evaluated. Cdk5 is known to phosphorylate several postsynaptic proteins including PSD-95, a major postsynapse scaffold protein (Morabito et al., 2004). Phosphorylation of PSD-95 by Cdk5 suppresses PSD-95 multimerization and results in reduced PSD-95-dependent clustering of the NMDA receptor subunit NR1 and voltage-gated potassium channel Kv1.4. Enlarged clusters of PSD-95 were observed in Cdk5-KO cortical neurons (Morabito et al., 2004).

5.3 Regulation of Dendritic Spine Remodeling A recent study implicated Cdk5 in dendrite spine remodeling (Fu et al., 2007). It is known that EphA4 is essential for the maintenance or elimination of dendritic spines (Murai et al., 2003) through the ephexin 1-RhoA pathway (Sahln et al., 2005). Cdk5-mediated phosphorylation of exphexin 1 was found to be required for ephrin A1/Eph4-mediated spine retraction (Fu et al., 2007). Spine morphology is also regulated by Cdk5-mediated WAVE1 phosphorylation via actin dynamics (Kim et al., 2006). The involvement of Cdk5 in spine regulation is also indicated by the increased density of dendritic spines in transgenic mice overexpressing p25 (Fisher et al., 2005).

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Role of Cdk5 in Higher Cognitive Function

6.1 Learning Memory Cdk5 phosphorylates the NMDA receptor subunit NR2A, and the inhibition of Cdk5 activity severely attenuates long-term potentiation (LTP) induction in CA1 hippocampal neurons (Li et al., 2001). Cdk5 activity is enhanced during associative learning and fear conditioning (Fischer et al., 2002), and the inhibition of Cdk5 activity impaired associative learning in mice (Fischer et al., 2002). Spatial learning was impaired in p35-KO mice (Ohshima et al., 2005). An electrophysiological study of hippocampal slices from p35-KO mice revealed an impaired LTD induction and a reduced LTP threshold (Ohshima et al., 2005). In addition, transient overexpression of p25 in transgenic mice enhanced the LTP and associative learning (Fisher et al., 2005). These data suggest that Cdk5 activity is critical for in vivo synaptic plasticity, learning, and memory. Conditional inactivation of Cdk5 in the adult mouse brain, however, indicates the opposite effect of reduced Cdk5 activity on LTP induction and associative learning (Hawasli et al., 2007). In the study by Hawasli et al. (2007), Cdk5 elimination resulted in an increase in NR2B at the synaptic surface, which in turn enhanced the hippocampal LTP and improved associative learning. This study revealed a new aspect of the Cdk5 function whereby NR2B degradation is facilitated by its direct interaction with the protease calpain. Because this phenotype did not reflect the loss of Cdk5-mediated phosphorylation of synaptic proteins, further studies are required to determine the effects of the lack of Cdk5-mediated phosphorylation on higher brain function.

6.2 Regulation of Dopaminergic Signaling and Drug Addiction Cdk5 is involved in dopamine signaling at both pre- and postsynapse. Inhibition of Cdk5 activity results in increased dopamine release in the striatum, indicating the presynaptic function of Cdk5 as a negative regulator of dopamine release (Chergui et al., 2004). Cdk5 also modulates the efficacy of postsynaptic dopamine signaling by phosphorylating DARPP-32 at Thr75, whereby DARPP-32 is converted into a PKA inhibitor (Bibb et al., 1999). Cdk5 was further demonstrated to participate in cocaine-induced changes in dopamine signaling. Chronic treatment of animals with cocaine has been shown to upregulate the expression of Cdk5 and p35 in the striatum through the induction of DfosB (Bibb et al., 2001). The increase in Cdk5 and p35 expression is accompanied by the upregulated phosphorylation of DARPP-32 at Thr75 and decreased phosphorylation of PKA-target proteins. Furthermore, Cdk5 inhibition potentiates the locomotor effects of chronic cocaine administration (Bibb et al., 2001). On the other hand, in transgenic mice overexpressing p35, elevation of Cdk5 activity attenuates cocaine-mediated dopamine signaling (Takahashi et al., 2005), indicating the inhibitory effects of Cdk5 on dopamine signaling. Cdk5 also affects dopamine signaling through the phosphorylation of tyrosine hydroxylase (TH), the rate-limiting enzyme in the synthesis of dopamine (Kansy et al., 2004; Moy and Tsai, 2004). Phosphorylation of TH by Cdk5 enhances TH activity and stability of the protein (Moy and Tsai, 2004).

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Cdk5 Function in the Peripheral Nervous System

7.1 Role of Cdk5 in Pain Signaling Cdk5 and p35 expression is observed in pain-sensing C-fiber neurons within the dorsal root and trigeminal ganglia (Pareek et al., 2006). Peripheral inflammation increases calpain activity that induces the cleavage of p35 to p25 and enhances Cdk5 activity (Pareek et al., 2006). These findings suggest the function of Cdk5 in pain signaling, and its role in pain sensing was examined using p35-KO mice and p35-overexpressing Tgp35 mice. When p35-KO mice were exposed to a hot water bath, their reflex actions were delayed; inversely, Tgp35 mice exhibited hyperalgesia (Pareek et al., 2006). Furthermore, C-fiber-specific conditional Cdk5-KO mice demonstrated hypoalgesia. Cdk5 phosphorylates TRPV1, and TRPV1-KO mice exhibit hypoalgesia, indicating that Cdk5-mediated TRPV1 phosphorylation is important for nociceptive signaling (Pareek et al., 2007).

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Cdk5 Function Outside the CNS

Previous studies have demonstrated the extraneuronal roles of Cdk5 (review in Rosales and Lee, 2006). In this section, we will describe only some of them, with a focus on muscle differentiation, secretory exocytosis in the pituitary gland, and insulin secretion in pancreatic b-cells.

8.1 Myogenesis and NMJ Formation The Cdk5 protein and Cdk5-associated kinase activity was detected during early myogenesis in mouse C2 cells (Lazaro et al., 1997). Cdk5 and p35 are expressed in embryonic muscles and are concentrated at the NMJ in adulthood (Fu et al., 2001). Cdk5-mediated phosphorylation of the NRG receptor ErbB3 is required for NRG-induced ErbB3 activation, the initiation of downstream signaling, and enhancement of AChR transcription (Fu et al., 2001). Cdk5 phosphorylates the transcription factor STAT3 (Fu et al., 2004). NRG enhances Cdk5-dependent STAT3 phosphorylation and the transcription of STAT3 target genes including those encoding c-fos and junB (Fu et al., 2004). Cdk5 also plays a role in AChR clustering. It has been shown to be required for the Ach-induced dispersion of AchR clustering (Lin et al., 2005). In the NMJ of the diaphragm in Cdk5-KO mice, the bandwidth of the AchR endplate is increased (Fu et al., 2005). In cultured myotubes from Cdk5-KO mice, AchR clusters are larger after agrin treatment than those from wild-type mice, indicating that Cdk5 may negatively regulate the AchR cluster size (Fu et al., 2005; Lin et al., 2005). This finding is consistent with those in the case of PSD-95 clustering in the postsynapse of the CNS, as described earlier.

8.2 Secretory Exocytosis A recent study indicates that Cdk5 is involved in hormonal exocytosis via the Rho family of small GTPases (Xin et al., 2004). Cdk5 phosphorylates Trio, a Rho GDP/GTP exchange factor (GEF), thereby increasing GEF activity and activating Rac. Inhibition of Cdk5 activity results in reduced Rac activation and reduced exocytosis because actin reorganization during exocytosis is controlled by the Cdk5-mediated activation of Trio (Xin et al., 2004).

8.3 Insulin Secretion in Pancreatic b-Cells Cdk5-mediated Munc18 phosphorylation in b-cells acts as a positive regulator of insulin exocytosis (Lilja et al., 2001). Under conditions of high glucose, however, the inhibition of Cdk5 activity enhanced insulin secretion (Wei et al., 2005a). Mice lacking p35 showed enhanced insulin secretion in response to high glucose challenge. On stimulation with high glucose, kinase inhibition increases Ca2+ influx across the L-type voltage-dependent Ca2+ channel (L-VDCC) in b-cells (Wei et al., 2005a).

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Deregulation of Cdk5 and Neurodegenerative Disorder

9.1 Production of p25 Changes Stability and Localization In cultured neurons, the calpain-mediated cleavage of p35 into p25 occurs under neurotoxic conditions such as exposure to oxidative stress and the presence of amyloid-b (Ab) peptide, glutamate, and ionomycin (Kusakawa et al., 2000; Lee et al., 2000). As mentioned earlier, p35 has a short half-life of 20–30 min. On phosphorylation by Cdk5, p35 is rapidly ubiquitinated and degraded by proteosomes. Proteolytic cleavage converts the p25 fragment into a substantially more stable form, and its half-life is extended by threefold to

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fivefold (Patrick et al., 1998; Patrick et al., 1999). Cleavage of p35 into p25 results in the removal of the myristoylation sequence from p35, thereby leading to the redistribution of Cdk5 activity from the membrane to the cytosol and nucleus (Patrick et al., 1999; Lee et al., 2000). p25 contains all the elements necessary for Cdk5 binding and its in vitro and in vivo activation. The generation of p25 is, therefore, likely to cause prolonged activation of mislocated Cdk5. Introduction of p25 into cultured neurons causes hyperphosphorylation of tau and induces apoptosis (Patrick et al., 1999). Inducible p25 transgenic mice that express p25 in the postnatal forebrain exhibit profound neurodegeneration, astrogliosis, and tau-associated pathology (Cruz et al., 2003). It was also demonstrated that transgenic mice coexpressing both mutant (P301L) human tau and p25 show accumulation of aggregated and hyperphosphorylated tau (Noble et al., 2003). These results indicate that p25 is neurotoxic in vivo.

9.2 p25 and AD Initial information regarding the involvement of Cdk5 in neurodegenerative disease was derived from the enhancement in p25 accumulation concomitant with tau hyperphosphorylation and elevated Cdk5 activity observed in the postmortem brains of AD patients (Patrick et al., 1999; Tseng et al., 2002). However, the analysis of postmortem AD brains by other groups revealed that the expression of p35, p25, and Cdk5 is comparable between control and AD brains (Yoo and Lubec, 2001; Taniguchi et al., 2001; Tandon et al., 2003). Hyperactivation of Cdk5 may regulate amyloid precursor protein (APP) processing. Cdk5 phosphorylates the intracellular domain of APP at Thr668 (Iijima et al., 2000), and this phosphorylation of APP has been suggested to increase in vitro APP processing and Ab production (Lee et al., 2003; Liu et al., 2003). Furthermore, the intracellular Ab levels are elevated in p25-inducible mice (Cruz et al., 2006). These investigators supposed that increased BACE1 levels due to aberrant axonal APP transport and increased APP phosphorylation at Thr668 because of elevated Cdk5 activity may account for the accumulation of Ab within neurons (Cruz et al., 2003). However, the involvement of Cdk5-mediated APP phosphorylation at Thr668 in APP processing is questioned by the analysis of knockin mice in which the Thr668 residue of APP is mutated to the nonphosphorylatable alanine residue (Sano et al., 2006) because the basal levels of Ab40 and 42 in the mouse brain remain unaltered in knockin mice. Other Cdk5 substrates such as presenilin-1 may be involved in APP processing (Lau et al., 2002). Even though the involvement of Cdk5 in APP processing has not been proved, Cdk5 may be involved in Ab-induced neuronal death in the brains of AD patients. Ab has been shown to induce an increase in the p25 levels in cultured neurons (Lee et al., 2000) as well as in transgenic mouse models (Otth et al., 2002; Oakley et al., 2006). Ab-induced cell death can be reduced by the inhibition of Ab-induced p25 elevation (Li et al., 2003b) or by Cdk5 inhibitors (Alvarez et al., 1999; Zheng et al., 2005). These results indicate the involvement of Cdk5 in the pathophysiology of AD.

9.3 Cdk5 in Ischemic Brain Injury p25 production was increased in a rat model of transient forebrain ischemia, and activated Cdk5 phosphorylates the NMDA receptor, causing excitotoxicity (Wang et al., 2003). Cdk5 inhibition by viral infection of Cdk5-DN form ameliorates neuronal cell death in the hippocampal CA1 region (Wang et al., 2003). Cdk5 also phosphorylates MEF2, a transcription factor that exerts prosurvival effects and inhibits the antiapoptotic function of MEF2. Phosphorylation of MEF2 by an increase in the Cdk5 level in the nucleus renders cortical neurons more susceptible to excitotoxicity and oxidative stress (Gong et al., 2003). Therefore, the deregulation of Cdk5 activity and its mislocation within the neurons exacerbates neuronal death not only by enhancing the cell death pathway but also by impairing prosurvival signaling.

9.4 Deregulation of Cdk5 and Parkinson’s Disease It has been shown that Cdk5 is hyperactivated and plays a major role in dopamine loss in the MPTP model of Parkinson’s disease (Smith et al., 2006). Activated Cdk5 induces dopaminergic loss by phosphorylating

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the cytoplasmic substrate Prx2 to inhibit its antioxidative ability (Qu et al., 2007) and by phosphorylating the nuclear target MEF2 to inhibit its prosurvival function (Smith et al., 2006). Cdk5 also phosphorylates Parkin (Avraham et al., 2007), a putative E3 ubiquitin-ligase (Shimura et al., 2000); mutations in this gene are the most frequent cause of familial Parkinson disease (Kitada et al., 1998). Parkin phosphorylation by Cdk5 decreases its E3 ubiquitin-ligase activity (Avraham et al., 2007). The significance of Parkin phosphorylation by Cdk5 in the pathophysiology of Parkinson’s disease needs to be studied in animal models and human patients.

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Cdk5 as a Drug Target

Accumulating experimental data on the roles of Cdk5 in neurodegenerative disorders indicate the potential effectiveness of Cdk5 inhibitors for the treatment of these disease conditions, including AD and Parkinson’s disease. Cdk5 inhibitors will also be useful for the treatment of type II diabetic patients (Wei et al., 2005a). Since selective inhibitors of Cdk5 remain to be evaluated in humans, their ability to safely inhibit Cdk5 is unknown. However, nonselective Cdk inhibitors with activities against Cdk5 have been profiled in preclinical toxicology and clinical studies (Fischer and Gisnella-Borradori, 2005). For example, phase 2 clinical trials are currently being conducted for roscovitine to test its activity against leukemia and lung and breast cancers. Basic data concerning potential safety issues such as mechanismbased toxicities, side effects, and off-target effects will be obtained when these drugs are used for the treatment of human patients with neurodegenerative disorders. Preclinical animal evaluation is currently being conducted for roscovitine and other Cdk inhibitors to test their activity against AD and stroke. The development of selective and brain-permeable inhibitors will lead to their successful application for the treatment of neurodegenerative disorders in humans. Further, a thorough understanding of the role of Cdk5 in the regulation of normal cognition is important for evaluating the effects of Cdk5 inhibitors on human disorders.

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Gilmore EC, Ohshima T, Goffinet AM, Kulkarni AB, Herrup K. 1998. Cyclin-dependent kinase 5-deficient mice demonstrate novel developmental arrest in cerebral cortex. J Neurosci 18: 6370-6377. Gleeson JG, Allen KM, Fox JW, Lamperti ED, Berkovic S, et al. 1998. Doublecortin, a brain-specific gene mutated in human X-linked lissencephaly and double cortex syndrome, encodes a putative signaling protein. Cell 92: 63-72. Gong X, Tang X, Wiedmann M, Wang X, Peng J, et al. 2003. Cdk5-mediated inhibition of the protective effects of transcription factor MEF2 in neurotoxicity-induced apoptosis. Neuron 38: 33-46. Gupt A, Sanada K, Miyamoto DT, Rovelstad S, Nadarajah B, et al. 2003. Layering defect in p35 deficiency is linked to improper neuronal-glial interaction in radial migration. Nat Neurosci 6: 1284-1291. Harada T, Morooka T, Ogawa S, Nishida E. 2001. ERK induces p35, a neuron-specific activator of Cdk5, through induction of Egr1. Nat Cell Biol 3: 453-459. Hawasli AH, Benavides DR, Nguyen C, Kansy JW, Hayashi K, et al. 2007. Cyclin-dependent kinase 5 governs learning and synaptic plasticity via control of NMDAR degradation. Nat Neurosci 10: 880-886. Hayashi F, Matsuura I, Kachi S, Maeda T, Yamamoto M, et al. 2000. Phosphorylation by cyclin-dependent protein kinase 5 of the regulatory subunit of retinal cGMP phosphodiesterase. II. Its role in the turnoff of phosphodiesterase in vivo. J Biol Chem 275: 32958-32965. Hellmich MR, Pant HC, Wada E, Battey JF. 1992. Neuronal cdc2-like kinase: A cdc2-related protein kinase with predominantly neuronal expression. Proc Natl Acad Sci USA 89: 10867-10871. Huang KX, Paudel HK. 2000. Ser67-phosphorylated inhibitor 1 is a potent protein phosphatase 1 inhibitor. Proc Natl Acad Sci USA 97: 5824-5829. Humber S, Dhavan R, Tsai LH. 2000. p39 activates cdk5 in neurons, and is associated with the actin cytoskeleton J Cell Sci 113: 975-983. Iijima K, Ando K, Takeda S, Satoh Y, Seki T, et al. 2000. Neuron-specific phosphorylation of Alzheimer’s beta-amyloid precursor protein by cyclin-dependent kinase 5. J Neurochem 75: 1085-1091. Kansy JW, Daubner SC, Nishi A, Sotogaku N, Lloyd MD, et al. 2004. Identification of tyrosine hydroxylase as a physiological substrate for Cdk5. J Neurochem 91: 374-384. Kawauchi T, Chihama K, Nabeshima Y, Hoshino M. 2006. Cdk5 phosphorylates and stabilizes p27kip1 contributing to actin organization and cortical neuronal migration. Nat Cell Biol 8: 17-26. Kesavapany S, Amin N, Zheng YL, Nijhara R, Jaffe H, et al. 2004. p35/cyclin-dependent kinase 5 phosphorylation

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of ras guanine nucleotide releasing factor 2 (RasGRF2) mediates Rac-dependent Extracellular Signal-regulated kinase 1/2 activity, altering RasGRF2 and microtubuleassociated protein 1b distribution in neurons. J Neurosci 24: 4421-4431. Kesavapany S, Lau KF, McLoughlin DM, Brownless J, Ackerley S, et al. 2001. p35/cdk5 binds and phosphorylates betacatenin and regulates beta-catenin/presenelin-1 interaction. Eur J Neurosci 13: 241-247. Keshvara L, Magdaleno S, Benhayon D, Curran T. 2002. Cyclin-dependent kinase 5 phosphorylates disabled 1 independently of Reelin signaling. J Neurosci 22: 4869-4877. Kim Y, Sung JY, Ceglia I, Lee KW, Ahn JH, et al. 2006. Phosphorylation of WAVE1 regulates actin polymerization and dendritic spine morphology. Nature 442: 814-817. Kitada T, Asakawa S, Hattori N, Matsumine H, Yamamura Y, et al. 1998. Mutations in the parkin. gene cause autosomal recessive juvenile parkinsonismNature 392: 605-608. Ko J, Humbert S, Brorson RT, Takahashi S, Kulkarni AB, et al. 2001. p35 and p39 are essential for cyclin-dependent kinase 5 function during neurodevelopment. J Neurosci 21: 6758-6771. Kobayashi S, Ishiguro K, Omori A, Takamatsu M, Arioka M, et al. 1993. A cdc2-related kinase PSSALRE/cdk5 is homologous with the 30 kDa subunit of tau protein kinase II, a proline-directed protein kinase associated with microtubule. FEBS Lett 335: 171-175. Kusakawa G, Saito T, Onuki R, Ishiguro K, Kishimoto T, et al. 2000. Calpain-dependent proteolytic cleavage of the p35 cyclin-dependent kinase 5 activator to p25. J Biol Chem 275: 17166-17172. Kwon YT, Gupta A, Zhou Y, Nikolic M, Tsai L-H. 2000. Regulation of the N-cadherin-mediated adhesion by the p35-Cdk5 kinase. Curr Biol 10: 363-372. Lau KF, Howlett DR, Kasavapany S, Standen CL, Dingwall C, et al. 2002. Cyclin-dependent kinase-5/p35 phosphorylates presenilin 1 to regulate carboxy-terminal fragment stability. Mol Cell Neurosci 20: 13-20. Lazaro JB, Kitzmann M, Poul MA, Vandromme M, Lamb NJ, et al. 1997. Cyclin dependent kinase 5, cdk5, is a positive regulator of myogenesis in mouse C2 cells. J Cell Sci 110: 1251-1260. Lee KY, Helbing CC, Choi KS, Johnston RN, Wang JH. 1997. Neuronal Cdc2-like kinase (Nclk) binds and phosphorylates the retinoblastoma protein. J Biol Chem 272: 5622-5626. Lee MS, Kao SC, Lemere CA, Xia W, Tseng HC, et al. 2003. APP processing is regulated by cytoplasmic phosphorylation. J Cell Biol 163: 83-95. Lee MS, Kwon YT, Li M, Peng J, Friedlander RM, et al. 2000. Neurotoxicity induces cleavage of p35 to p25 by calpain. Nature 405: 360-364.

Lee SY, Wenk MR, Kim Y, Nairn AC, De Camili P. 2004. Regulation of synaptojanin 1 by cyclin-dependent kinase 5 at synapses. Proc Natl Acad Sci USA 101: 546-551. Lew J, Beaudette K, Litwin CM, Wang JH. 1992. Purification and characterization of a novel proline-directed protein kinase from bovine brain. J Biol Chem 267: 13383-13390. Lew J, Huang QQ, Qi Z, Winkfein RJ, Aebersold R, et al. 1994. A brain-specific activator of cyclin-dependent kinase 5. Nature 371: 423-425. Li BS, Ma W, Jaffe H, Zheng Y, Takahashi S, et al. 2003a. Cyclin-dependent kinase-5 is involved in neuregulindependent activation of phosphatidylinositol 3-kinase and Akt activity mediating neuronal survival. J Biol Chem 278: 35702-35709. Li BS, Sun MK, Zhang L, Takahashi S, Ma W, et al. 2001. Regulation of NMDA receptors by cyclin-dependent kinase-5. Proc Natl Acd Sci USA 98: 12742-12747. Li BS, Zhang L, Gu J, Amin ND, Pant HC. 2000. Integrin alpha(1) beta(1)-mediated activation of cyclin-dependent kinase 5 activity is involved in neurite outgrowth and human neurofilament protein H Lys-Ser-Pro tail domain phosphorylation. J Neurosci 15: 6055-6062. Li BS, Zhang L, Takahashi S, Ma W, Jaffe H, et al. 2002. Cyclin-dependent kinase 5 prevents neuronal apoptosis by negative regulation of c-Jun N-terminal kinase 3. EMBO J 21: 324-333. Li A, Faibushevich G, Turunen BJ, Yoon SO, Georg G, et al. 2003b. Stabilization of the cyclin-dependent kinase 5 activator, p35, by paclitaxel decreases beta-amyloid toxicity in cortical neurons. J Neurochem 84: 347-362. Lilja L, Yang SN, Webb DL, Juntti-Berggren L, Berggren PO, et al. 2001. Cyclin-dependent kinase 5 promotes insulin exocytosis. J Biol Chem 276: 34199-34205. Lim AC, Hou Z, Goh CP, Qi RZ. 2004. Protein kinase CK2 is an inhibitor of the neuronal Cdk5 kinase. J Biol Chem 279: 46668-46673. Lin W, Dominguez B, Yang J, Aryal P, Brandon EP, et al. 2005. Neurotransmitter acetylcholine negatively regulates neuromuscular synapse formation by a Cdk5-dependent mechanism. Neuron 46: 569-579. Liu F, Su Y, Li B, Zhou Y, Ryder J, et al. 2003. Regulation of amyloid precursor protein (APP) phosphorylation and processing by p35/Cdk5 and p25/Cdk5. FEBS Lett 547: 193-196. Matsubara M, Kusubata M, Ishiguro K, Uchida T, Titani K, et al. 1996. Site-specific phosphorylation of synapsin I by mitogen-activated protein kinase and Cdk5 and its effects on physiological functions. J Biol Chem 271: 21108-21113. Matsuura I, Bondarrenko VA, Maeda T, Kachi S, Yamazaki M, et al. 2000. Phosphorylation by cyclin-dependent protein

Cyclin-dependent kinase 5 kinase 5 of the regulatory subunit of retinal cGMP phosphodiesterase. I. Identification of the kinase and its role in the turnoff of phosphodiesterase in vitro. J Biol Chem 275: 32950-32957. Meyerson M, Enders GH, Wu CL, Su LK, Gorka C, et al. 1992. A family of human CDC2-related protein kinases. EMBO J 11: 2909-2917. Morabito MA, Sheng M, Tsai LH. 2004. Cyclin-dependent kinase 5 phosphorylates the N-terminal domain of the postsynaptic density protein PSD-95 in neurons. J Neurosci 24: 865-876. Moreno S, Nurse P. 1990. Substrates for p34cdc2: In vivo veritas? Cell 61: 549-551. Moy LY, Tsai LH. 2004. Cyclin-dependent kinase 5 phosphorylates serine 31 of tyrosine hydroxylase and regulates its stability. J Biol Chem 279: 54487-54493. Murai KK, Nguyen LN, Irie F, Yamaguchi Y, Pasquale EB. 2003. Control of hippocampal dendritic spine morphology through ephrin-A3/EphA4 signaling. Nat Neurosci 6: 153160. Niethammer M, Smith DS, Ayala R, Peng J, Ko J, et al. 2000. NUDEL is a novel Cdk5 substrate that associates with LIS1 and cytoplasmic dynein. Neuron 28: 697-711. Nikolic M, Chou MM, Lu W, Mayer BJ, Tsai L-H. 1998. The p35/Cdk5 kinase is a neuron-specific Rac effector that inhibits Pak1 activity. Nature 395: 194-198. Nikolic M, Dudek H, Kwon YT, Ramos YF, Tsai L-H. 1996. The cdk5/p35 kinase is essential for neurite outgrowth during neuronal differentiation. Genes Dev 10: 816-825. Noble W, Olm V, Takata K, Casey E, Mary O, et al. 2003. Cdk5 is a key factor in tau aggregation and tangle formation in vivo. Neuron 38: 555-565. Oakley H, Cole SL, Logan S, Maus E, Shao P, et al. 2006. Intraneuronal beta-amyloid aggregates, neurodegeneration, and neuron loss in transgenic mice with five familial Alzheimer’s disease mutations: Potential factors in amyloid plaque formation. J Neurosci 26: 10129-10140. Ohshima T, Hirasawa M, Tabata H, Mutoh T, Adachi T, et al. 2007b. Cdk5 is required for multipolar-to-bipolar transition during radial neuronal migration and proper dendrite development of pyramidal neurons in the cerebral cortex. Development 134: 2273-2282. Ohshima T, Mikoshiba K. 2002. Reelin signaling and Cdk5 in the control of neuronal positioning. Mol Neurobiol 26: 153-166. Ohshima T, Ogawa M, Takeuchi K, Takahashi S, Kulkarni AB, et al. 2002. Cyclin-dependent kinase 5/p35 contributes synergistically with Reelin/Dab1 to the positioning of facial branchiomoter and inferior olive neurons in the developing mouse hindbrain. J Neurosci 22: 4036-4044. Ohshima T, Ogawa M, Veeranna HM, Longenecker G, Ishiguro K, et al. 2001. Synergistic contributions of

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Takahashi S, Ohshima T, Cho A, Sreenth T, Iadaroa MJ, et al. 2005. Increased activity of cyclin-dependent kinase 5 leads to attenuation of cocaine-mediated dopamine signaling. Proc Natl Acad Sci USA 102: 1737-1742. Tan TC, Valova VA, Malladi CS, Graham ME, Berven LA, et al. 2003. Cdk5 is essential for synaptic vesicle endocytosis. Nat Cell Biol 5: 701-710. Tanaka T, Serneo FF, Tseng HC, Kulkarni AB, Tsai L-H, et al. 2004. Cdk5 phosphorylation of doublecortin ser297 regulates its effect on neuronal migration. Neuron 41: 215-227. Tandon A, Yu H, Wang L, Rogaeva E, Sato C, et al. 2003. Brain levels of CDK5 activator p25 are not increased in Alzheimer’s or other neurodegenerative diseases with neurofibrillary tangles. J Neurochem 86: 572-581. Tang D, Yeung J, Lee KY, Matsushita M, Matsui H, et al. 1995. An isoform of the neuronal cyclin-dependent kinase 5 (Cdk5) activator. J Biol Chem 270: 26897-26903. Taniguchi S, Fujita Y, Hayashi S, Kakita A, Takahashi H, et al. 2001. Calpain-mediated degradation of p35 to p25 in postmortem human and rat brains. FEBS Lett 489: 46-50. Tarricone C, Dhavan R, Peng J, Areces LB, Tsai LH, et al. 2001. Structure and regulation of the CDK5-p25(nck5a) complex. Mol Cell 8: 657-669. Tokuoka H, Saito T, Yorifuji H, Wei F, Kishimoto T, et al. 2000. Brain-derived neurotrophic factor-induced phosphorylation of neurofilament-H subunit in primary cultures of embryonic rat cortical neurons. J Cell Sci 113: 1059-1068. Tomizawa K, Ohta J, Matsushita M, Moriwaki A, Li ST, et al. 2002. Cdk5/p35 regulates neurotransmitter release through phosphorylation and downregulation of P/Q-type voltage-dependent calcium channel activity. J Neurosci 22: 2590-2597. Tomizawa K, Sunada S, Lu YF, Oda Y, Kinuta M, et al. 2003. Cophosphorylation of amphiphysin I and dynamin I by Cdk5 regulates clathrin-mediated endocytosis of synaptic vesicles. J Cell Biol 163: 813-824. Tsai L-H, Delalle I, Caviness VS JrChae T, Harlow E. 1994. p35 is a neural-specific regulatory subunit of cyclin-dependent kinase 5. Nature 371: 419-423. Tsai L-H, Takahashi T, Caviness VS JrHarlow E. 1993. Activity and expression pattern of cyclin-dependent kinase 5 in the embryonic mouse nervous system. Development 119: 1029-1040. Tseng HC, Zhou Y, Shen Y, Tsai LH. 2002. A survey of Cdk5 activator p35 and p25 levels in Alzheimer’s disease brains. FEBS Lett 523: 58-62. Uchida Y, Ohshima T, Sasaki Y, Suzuki H, Yanai S, et al. 2005. Semaphorin3A signalling is mediated via sequential Cdk5 and GSK3b phosphorylation of CRMP2: Implication of

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N. Maeda

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204

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PTP Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204 PTP Family Members . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204 Structures of RPTPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 206

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Regulation of the Catalytic Activity of RPTPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 206 Regulation of RPTP Function by Dimerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 206 Regulation of RPTP Function by Reversible Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 208

4 4.1 4.2 4.2.1 4.2.2 4.2.3 4.2.4 4.3 4.3.1 4.3.2 4.3.3 4.3.4 4.3.5

Functions of RPTPs in the Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 Proteoglycans in the Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 PTPz/Phosphacan and Neural Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 Structure of PTPz/Phosphacan . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210 The Binding of Pleiotrophin/Midkine with Phosphacan . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 212 Pleiotrophin Is a Ligand of PTPz . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 213 Pleiotrophin Induces Oligomerization of PTPz . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214 DLAR and Neural Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214 Drosophila RPTPs and Glycosaminoglycans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 214 RPTP and Motor Axon Guidance in Drosophila . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 215 DLAR and Syndecan in Motor Axon Guidance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 215 Interaction of DLAR with Syndecan and Dallylike . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 215 DLAR, Syndecan, and Dallylike in Synaptogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 216

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Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217

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Abstract: Receptor-like protein tyrosine phosphatases are composed of an extracellular domain, a transmembrane segment and one or two tyrosine phosphatase domains. Their extracellular regions show several domain structures including immunoglobulin-like, fibronectin type III-like, carbonic anhydrase-like, and meprin/A5/m domains. The regulatory mechanisms of their tyrosine phosphatase activities are still unclear. However, it seems that ligand-induced inactivation or activation and reversible oxidation of catalytic cysteine residue are the major signaling mechanisms. Receptor-like protein tyrosine phosphatases are involved in various cell–cell and cell–extracellular matrix interactions, and accumulating evidence has revealed that they play important roles in the nervous system, such as axon guidance, neurite outgrowth, neuronal migration, and learning processes. Recent studies indicated that proteoglycans are essential components of the signal transduction pathways of several receptor-like protein tyrosine phosphatases. Chondroitin sulfate and heparan sulfate regulate ligand binding to PTPz and DLAR, respectively, and structural variations of these glycosaminoglycans may be involved in the determination of the binding affinity. List of Abbreviations: ECM, extracellular matrix; EGFR, epidermal growth factor receptor; H2O2, hydrogen peroxide; MAM, meprin/A5/m; PTK, protein tyrosine kinases; PTP, protein tyrosine phosphatases; ROS, reactive oxygen species; RPTP, receptor-like PTPs

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Introduction

Tyrosine phosphorylation is utilized exclusively in multicellular eukaryotes and is rare in prokaryotes, although protein phosphorylation itself is ubiquitously observed (Alonso et al., 2004). This suggests that tyrosine phosphorylation is necessary for complex cell–cell and cell–extracellular matrix (ECM) interactions that regulate cell proliferation, differentiation, migration, adhesion, and morphogenesis in multicellular organisms. Accordingly, the development and the functioning of nervous system should strongly depend on tyrosine phosphorylation processes, because nervous system is the most complex tissue in the organism. The tyrosine phosphorylation levels of intracellular proteins are determined by the coordinated activities of protein tyrosine kinases (PTKs) and protein tyrosine phosphatases (PTPs). Although much knowledge about PTK functions in the nervous system has been accumulated, that about PTPs is still scarce. However, recent studies have begun to elucidate the essential roles of PTPs in the development and function of nervous tissues. On the other hand, it has been recognized that proteoglycans play pivotal roles in the development and maintenance of the nervous system as major components of the cell surface and ECM. Proteoglycans bind with various proteins such as growth factors and receptors through their core protein and glycosaminoglycan portions. Glycosaminoglycans, especially chondroitin sulfate and heparan sulfate, display enormous structural heterogeneity. The binding between proteoglycans and many proteins is dependent on the structure of glycosaminoglycans, and structurally distinct glycosaminoglycans exhibit different affinities for various proteins. Thus, glycosaminoglycans are believed to enable diversification of many molecular interactions that are necessary for the development of complex tissues. Recent studies revealed that proteoglycans are an essential part of the signaling pathways of some PTPs. In this chapter, I will discuss the structure and function of receptor-like PTPs (RPTPs), and their functional relationship with proteoglycans.

2

PTP Family

2.1 PTP Family Members PTPs are a large family of enzymes defined by an active-site signature motif, HCX5R, in which the cysteine residue functions as a nucleophile that attacks the substrate phosphate. PTP family members are generally classified into two groups: the classical phosphotyrosine-specific phosphatases and the dual-specificity

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phosphatases, which may also dephosphorylate serine and threonine residues and lipid substrates (Alonso et al., 2004; Tonks, 2006; Tiganis and Bennett, 2007). The classical phosphotyrosine-specific phosphatases are further grouped into the RPTPs and the cytoplasmic nonreceptor PTPs. An exhaustive search of the human genome revealed that there are 107 human PTP family genes (Alonso et al., 2004). Among them, 38 genes are classical PTPs, of which 21 genes encode RPTPs (> Figure 11-1). Regulation of the gene

. Figure 11-1 Schematic representation of RPTPs. Based on the domain structures, vertebrate RPTPs are classified into eight groups. The structures depicted here are only schematic and do not correspond to the exact structure of each RPTP. The names of RPTPs are extremely confusing because many of RPTPs are called by multiple names. Furthermore, even distinct RPTPs are sometimes called by the same or similar names. In this figure, only the most commonly used names are shown (see the reference by Andersen et al. (2001) for a complete list of synonyms). Names of Drosophila RPTPs are shown in parentheses

expression by the use of alternative promoters and alternative mRNA splicing and posttranslational modifications add further complexity to this diverse enzyme family (Tonks, 2006). Especially, the extracellular regions of several RPTPs are highly glycosylated, modifying the ligand–receptor interaction (see later). Such enormous diversity of PTPs should contribute to the complex cell–cell interactions that are essential for the development and maintenance of the multicellular organism.

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2.2 Structures of RPTPs RPTPs are composed of an extracellular region, a transmembrane segment, and one or two tyrosine phosphatase domains (D1 and D2 domains) (> Figure 11-1). The extracellular portions include immunoglobulin, fibronectin type III, meprin/A5/m (MAM), and carbonic anhydrase domains, and are thought to interact with extracellular ligands regulating the cell–cell and cell–ECM interaction. Based on the extracellular domain structures and the sequence homology of catalytic domains, the RPTPs are classified into eight subgroups (Andersen et al., 2001; Johnson and Van Vactor, 2003) (> Figure 11-1). The only type I/VI RPTP is CD45, which is highly expressed on the surface of hematopoietic cells. The extracellular domain of this PTP contains fibronectin type III domains and is highly glycosylated. The type IIA subfamily are characterized by the cell adhesion molecule-like extracellular domains consisting of multiple immunoglobulin and fibronectin type III domains. Vertebrate LAR, PTPs, PTPd, and Drosophila DLAR and DPTP69D belong to this subfamily. The type IIB subfamily have an Nterminal MAM domain, an immunoglobulin domain, and multiple fibronectin type III domains in the extracellular region. PTPm, PTPk, PTPl, and PTPr belong to this subfamily. The type III subfamily is characterized by multiple fibronectin type III domains in the extracellular region and a single intracellular phosphatase domain. Vertebrate PTPb, DEP1, SAP1, GLEPP1, and PTPS31, and Drosophila DPTP99A, DPTP10D, DPTP4E, and DPTP52F belong to this family. Among these, DPTP99A is an exception in that it has two cytoplasmic phosphatase domains. The type IV subfamily (PTPa and PTPe) have very short extracellular regions that may be heavily glycosylated. The type V subfamily (PTPg and PTPz) is characterized by an N-terminal carbonic anhydrase-like domain and a fibronectin type III domain. PTPz is quite unique because it is synthesized as chondroitin sulfate proteoglycan (see later). The type VII subfamily (PCPTP1 and STEP) is characterized by a short extracellular region, a cytoplasmic kinase-interaction motif, and one phosphatase domain. The extracellular region of type VIII subfamily members contains an RDGS-adhesion recognition motif, but the members of this family (IA2 and IA2b) are considered to be catalytically inactive. Most of the PTP activity of RPTPs resides in the membrane-proximal D1 domain, except in the case of PTPa, in which both D1 and D2 domains have PTP activity (Groen et al., 2005). In spite of the absence of phosphatase activity, the D2 domain is considered to be important for determination of the substrate specificity and gathering of the downstream signaling molecules (Streuli et al., 1990; Debant et al., 1996).

3

Regulation of the Catalytic Activity of RPTPs

Due to the transmembrane receptor-like structure, it has been believed that extracellular ligands regulate the PTP activities of RPTPs. In fact, many homophilic interactions and extracellular binding partners have been found for RPTPs; however, these were found to regulate the catalytic activity in only a few cases (> Table 11-1). Several models describing how RPTPs transduce extracellular signals into the cytoplasm have been proposed, although many of them are still hypothetical (> Figure 11-2).

3.1 Regulation of RPTP Function by Dimerization Initially, a dimerization-based model was worked out from the elucidation of the crystal structure of the D1 domain of PTPa and the subsequent biochemical analysis of an epidermal growth factor receptor (EGFR)CD45 chimeric molecule (Bilwes et al., 1996; Majeti et al., 1998). In the crystal structure, the D1 domains of PTPa formed dimers, in which the N-terminal helix-turn-helix segment (inhibitory wedge) of one domain inserts into the catalytic cleft of the other domain (Bilwes et al., 1996). Using an EGFR-CD45 chimera composed of the extracellular domain of EGFR and the intracellular segment of CD45, Majeti et al. (1998) showed that ligand-induced dimerization inhibited the function of this RPTP in T cell signal transduction, which requires intact PTP activity. From these observations, it was proposed that reciprocal active-site blockage by PTP dimerization is a general signal transduction mechanism of RPTPs. So, if RPTP is present

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. Table 11-1 Binding partners of RPTPs Receptor-type Type I/VI Type IIA

PTP CD45 LAR

PTPσ PTPδ DLAR Type IIB

Type III Type IV Type V

PTPμ PTPκ PTPλ DEP1 PTPα PTPζ

Binding proteins galectin-1 Homophilic binding (small ectodomain isoform) Laminin-Nidogen complex Heparan sulfate proteoglycan Nucleolin Homophilic binding Syndecan Dallylike Homophilic binding Homophilic binding Homophilic binding Matrigel Contactin Pleiotrophin midkine, FGF-2 F3/contactin, TAG-1, Nr-CAM Tenascin, N-CAM, L1/Ng-CAM

PTP activity Inactivation

Inactivation

Activation Inactivation

References Walzel H, et al. (1999) Yang T, et al. (2003) O’Grady P, et al. (1998) Aricescu AR (2002) Alete DE (2006) Wang J and Bixby JL (1999) Fox AN, et al. (2005) Johnson KG, et al. (2006) Brady-Kalnay SM (1994) Sap J, et al. (1994) Cheng J (1997) Sorby M (2001) Zeng L (1999) Maeda N, et al. (1996); Meng K, et al. (2000) Maeda N (2007) for review

as a monomer on the cell surface in the unstimulated state, ligand-induced dimerization will inactivate the PTP activity (> Figure 11-2b). On the other hand, if the RPTP constitutively forms dimer on the plasma membrane, ligands may dissociate the dimer and activate the PTP activity (> Figure 11-2c). Homophilic and heterophilic interaction of RPTPs in trans with RPTPs or other cell surface molecules on the adjacent cells may also influence the dimerization process (> Figure 11-2a). Jiang et al. (1999, 2000) showed that PTPa dimerizes constitutively on the living cell surface and is inactivated, suggesting that ligand-induced activation works in this case. Contactin binds with PTPa in a cis configuration and forms a neuronal receptor complex coupled with fyn tyrosine kinase (Zeng et al., 1999). Such cis interaction may also regulate the dimerization process. Sorby et al. (2001) reported that Matrigel, a commercially available mixture of ECM components prepared from Englebreth-Holm-Swarm mouse sarcoma, activated the PTP activity of DEP-1. Recent small-angle X-ray scattering analysis indicated that the catalytic domain of DEP-1 (PTPZ) forms a dimer that is closely related to the autoinhibitory D1 dimer of PTPa (Matozo et al., 2007). Thus, some ECM components in Matrigel might act as ligands that disrupt the DEP1 dimer and activate its PTP activity. However, it should be noted that the D1–D1 interaction of DEP1 is concentration dependent with a dissociation constant of 21.6 mM (Matozo et al., 2007). This suggests that DEP-1 may be present principally as an active monomer in the cells expressing low levels of this enzyme and be inactivated by ligand-induced dimerization. Furthermore, ligand-induced dimerization and the following inactivation of PTP activity was proposed as the signal transduction mechanism of CD45 (Walzel et al., 1999) and PTPz (see later). A variation of the dimerization-based model was proposed for type IIA LAR family members (Wallace et al., 1998). PTPs-D1 binds with PTPd-D2, which leads to the inhibition of the catalytic activity of PTPsD1, suggesting that receptor heterodimerization works as another signaling mechanism. However, a dimerization-based mechanism may not be a general feature of RPTPs, because the crystal structure of D1 of PTPm showed that the catalytic sites are not hindered and adopt an open conformation in the D1 dimer (Hoffmann et al., 1997). In this case, intramolecular interactions between the juxtamembrane domain and the PTP domains may regulate the PTP activity (Feiken et al., 2000).

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. Figure 11-2 Models for the regulation of RPTP function. (a) RPTPs may interact with proteins on adjacent cells in a homophilic or heterophilic manner. It is unclear whether such interactions influence the PTP activity. (b) Extracellular ligands may induce dimerization of RPTPs, leading to the inactivation of PTP activity. (c) When RPTPs spontaneously dimerize, extracellular ligands may disrupt the dimer leading to the activation of PTP activity

3.2 Regulation of RPTP Function by Reversible Oxidation Recently, reversible oxidation of catalytic sites by reactive oxygen species (ROS) was proposed as a signal transduction mechanism of RPTPs (Meng et al., 2002; Tonks, 2005). Although ROS have long been regarded as harmful ‘‘accidental’’ metabolic byproducts, recent studies suggest that cells produce ROS in a spatially restricted manner, and that these ROS work as a local signaling mediator (Lambeth, 2004; Terada, 2006). NADPH oxidase family members catalyze the transfer of an electron from NADPH to molecular oxygen to generate superoxide (O2 ), which in turn yields other ROS, such as hydrogen peroxide (H2O2).

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ROS are generated in various tissues in response to growth factors, cytokines, and calcium signals by the activation of NADPH oxidases through phosphorylation of the regulatory proteins and Rac activation (Lambeth, 2004). ROS oxidize specific proteins reversibly, and it is now considered that such reversible oxidation regulates various signaling pathways in a manner analogous to reversible protein phosphorylation (Meng et al., 2002). Specific cysteine residues in some types of proteins are easily oxidized by ROS and converted to the sulfenic acid form (–SOH). PTPs are one such type of proteins. As described earlier, the cysteine residue in the active center of PTPs works as a strong nucleophile. This is because the pKa value of the sulfhydryl group of this cysteine residue is very low compared with those of typical cysteine residues (pKa = 8.5), which also makes it susceptible to oxidation (Meng et al., 2002). The thiolate anion (–S ) in the active site cysteine residues can be oxidized to either sulfenic acid (–SOH), sulfinic acid (–SO2H), or sulfonic acid (–SO3H). The oxidized forms of cysteine lose nucleophilic activity, and therefore PTPs become inactive (> Figure 11-3). Although the sulfenic acid form of PTP is easily reduced back to the active form by cellular

. Figure 11-3 Regulation of RPTPs by reversible oxidation. Reactive oxygen species (ROS) are generated in various cells after stimulation with growth factors. RPTPs may be transiently inactivated by ROS and strengthen the signaling of protein tyrosine kinases by growth factor stimulation, after which RPTP may be reactivated and terminate the growth factor signaling. The thiolate anion (–S ) in the active-site cysteine residue can be oxidized to either sulfenic acid (–SOH), sulfinic acid (–SO2H), or sulfonic acid (–SO3H). Although the sulfenic acid form of PTP is easily reduced back to the active form, oxidation to sulfinic acid or sulfonic acid is irreversible

thiols such as glutathione and thioredoxin, oxidation to sulfinic or sulfonic acid is usually irreversible (Tonks, 2005). Thus, when pervanadate, a reagent that is often used as a PTP inhibitor, irreversibly oxidizes PTPs to the sulfonic acid form (Persson et al., 2004), reversible oxidization to the sulfenic acid form by H2O2 can be a physiological regulatory mechanism of PTP activity. A series of studies on PTPa (Blanchetot et al., 2002; Persson et al., 2004; Van der Wijk et al., 2004; Groen et al., 2005) indicated that the D2 domain is preferentially oxidized compared with the D1 domain, suggesting that the D2 domain of PTPa may function as a redox sensor. Treatment of the cells expressing PTPa with H2O2 induced oxidation of catalytic cysteine in the D2 domain, which led to stabilization of the inactive PTPa dimer by the formation of an intermolecular disulfide bond (Van der Wijk, 2004). Removal of oxidative stress first reduced the disulfide bonds, and PTPa slowly refolded to the active conformation. Thus, reversible oxidation of the catalytic cysteine residue may modulate the dimerization-based signaling mechanism of RPTPs. However, further studies will be needed to know whether this is a general mechanism applicable to other RPTPs. In fact, it was reported that the D1 domain of LAR was readily oxidized, suggesting that LAR is directly inactivated by ROS-induced oxidization of the catalytic cysteine residue in the D1 domain (Groen et al., 2005). On the other hand, the D1 domain of PTPm was not readily oxidized like that of PTPa (Groen et al., 2005). These findings suggest that RPTPs are differentially regulated by reversible oxidation.

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Functions of RPTPs in the Nervous System

Recent studies have demonstrated the functional importance of RPTPs in the nervous system, and representative reports are listed in > Table 11-2. As expected from the fact that tyrosine phosphorylation is involved in the diverse physiological processes in the nervous system, RPTPs play a variety of roles, including regulation of neurite outgrowth, axon guidance, neuronal migration, synaptogenesis, ion channels and NMDA receptor functions, motor coordination, and learning processes. Because there is not enough space to describe each of these functions, I will focus on PTPz and DLAR, especially discussing the involvement of proteoglycans in the signal transduction of these RPTPs.

4.1 Proteoglycans in the Nervous System Proteoglycans are complex glycoconjugates composed of a core protein and one or more glycosaminoglycan chains that are covalently attached to it. Glycosaminoglycans are classified into hyaluronic acid, chondroitin sulfate, heparan sulfate, and keratan sulfate (> Figure 11-4). Although hyaluronic acid is a simple polysaccharide without sulfation, chondroitin sulfate and heparan sulfate are highly sulfated and display enormous structural heterogeneity because of the differential sulfation patterns. It is believed that various proteins, including growth factors and ECM molecules, read the difference in the sulfation patterns of these glycosaminoglycans. That is, glycosaminoglycans with different sulfation patterns are considered to exhibit different affinities for such proteins, and therefore differentially regulate their functions. Lectican (hyalectan) family members (aggrecan, versican, neurocan, and brevican), NG2, neuroglycan C, and PTPz/phosphacan are the major chondroitin sulfate proteoglycans in the nervous system, while transmembrane syndecan and GPI-anchored glypican family members are the major heparan sulfate proteoglycans in the brain. Although syndecan family members are usually regarded as heparan sulfate proteoglycans, they may also contain chondroitin sulfate. Hyaluronic acid is synthesized as free long polysaccharides and is not present as proteoglycans. Instead, lectican family members bind noncovalently with hyaluronic acid, forming large complexes that are stabilized by link proteins. These proteoglycans are the major components of the ECM and cell surface of the nervous system, and play important roles in cell proliferation, migration, differentiation, adhesion, and morphogenesis (Bandtlow and Zimmermann, 2000). Although it is well known that heparan sulfate proteoglycans modulate the signaling of many heparin-binding growth factors such as FGF-2, little is known about the proteoglycan-mediated signal transduction mechanism that is involved in nervous system development. However, recent studies have indicated that proteoglycans are deeply involved in the signal transduction of RPTPs.

4.2 PTPz/Phosphacan and Neural Functions 4.2.1 Structure of PTPz/Phosphacan PTPz is a quite unique RPTP because it is synthesized as a chondroitin sulfate proteoglycan. The extracellular domain of this RPTP consists of an N-terminal carbonic anhydase-like domain, a fibronectin type III domain, and a large serine-glycine-rich region (> Figure 11-1). There are three major splicing variants: (1) the full-length receptor form (PTPz-A), (2) the short receptor form, in which most of the serine–glycine-rich region is deleted (PTPz-B), and (3) the secreted form, which corresponds to the extracellular region of PTPz-A (phosphacan). Phosphacan, as well as neurocan, is the major soluble chondroitin sulfate proteoglycan in the brain. Although phosphacan and PTPz-A are constantly synthesized as chondroitin sulfate proteoglycans, PTPz-B is a so-called part-time proteoglycan, and a nonproteoglycan form is detected in the rat brain at the early embryonic stages (Nishiwaki et al., 1998). The structure of chondroitin sulfate chains on phosphacan dynamically changes during development of the brain (Maeda et al., 2003), suggesting that chondroitin sulfate plays a role in important regulatory functions.

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. Table 11-2 Functions of RPTPs in the nervous system Receptortype Type I/VI Type IIA

PTP CD45 LAR

PTPσ

PTPδ

DLAR DPTP69D Type IIB

PTPμ PTPκ

Type III

GLEPP1

Type IV

DEP1 DPTP99A DPTP10D DPTP52F PTPα

PTPε Type V

PTPγ PTPζ

Reported functions Regulation of microglial activation (1) Spatial learning (2) Regulation of adult dentate gyrus neurogenesis (3) Development and target projection of basal forebrain cholinergic neurons (4) Development and maintenance of exitatory synapses in hippocampal neurons (1) Motor axon growth and guidance (2) Neuroendocrine dysplasia in KO mice (1) Motor axon growth and guidance (2) Learning and hippocampal LTP (3) Neurite promoting activity in the extracellular domain (1) Motor axon guidance and targeting (2) Targeting of photoreceptor neurons (1) Motor axon guidance and retinal axon targeting (2) Retinal axon targeting Retino-tectal projection Neurite promoting activity in the extaracellular domain (1) Motor axon growth and guidance (2) Neurite inhibitory guidance cue for retinal neurons Somatostatin inhibition of glioma proliferation Motor axon guidance Axon guidance across the CNS midline CNS and motor axon guidance (1) Spatial learning (2) Radial neuronal migration in hippocampus (3) Retinal development (1) Myelination of sciatic nerve (2) Kv channel regulation in Schwann cells Motor deficit in KO mice (1) Migration of cortical neurons and gliomas (2) Neurite extension

Type VII

PCPTP STEP

(3) Spatial learning (4) Oligodendrocyte survival and recovery from demyelinating disease (5) Regulation of sodium channels Motor coordination Regulation of NMDA receptor functions

References Tan J, et al. (2000) Kolkman MJM, et al. (2004) Bernabeu R, et al. (2006) Yeo TT, et al. (1997); Van Lieshout EMM, et al. (2001) Dunah AW, et al. (2005) Stepanek L, et al. (2005) Elchebly M, et al. (1999); Wallace MJ (1999) Stepanek L, et al. (2005) Uetani N, et al. (2000) Uetani N, et al. (2006) Krueger NX, et al. (1996); Clandinin TR, et al. (2001) Desai CJ, et al. (1996); Garrity PA, et al. (1999) Burden-Gulley SM, et al. (2002) Drosopoulos NE, et al. (1999) Stepanek L, et al. (2005) Stepanek L, et al. (2001) Massa A, et al. (2004) Desai CJ, et al. (1996) Sun Q, et al. (2000) Schindelholz B, et al. (2001) Skelton MR, et al. (2003) Petrone A, et al. (2003) van der Sar AM, et al. (2002) Tiran Z, et al. (2006) Tiran Z, et al. (2006) Lamprianou S, et al. (2006) Maeda N, et al. (1998); Muller S, et al. (2003) Maeda N, et al. (1996); Tanaka M, et al. (2003) Niisato K, et al. (2005) Harroch S (2002) Ratcliffe C, et al. (2000) Chirivi RG, et al. (2007) Braithwaite SP, et al. (2006); Paul S, et al. (2003); Pelkey KA (2002)

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. Figure 11-4 Disaccharide units of glycosaminoglycans. Hyaluronic acid is composed of repeating disaccharide units of glucuronic acid (GlcA) and N-acetylglucosamine (GlcNAc) and is not sulfated. Chondroitin sulfate is composed of repeating disaccharide units of GlcA and N-acetylgalactosamine (GalNAc). GlcA residues are sometimes C5 epimerized to iduronic acid (arrow). Potential sulfation sites are shown. Heparan sulfate is composed of repeating disaccharide units of GlcA and GlcNAc. GlcA residues may be C5 epimerized (arrow). Potential sites for sulfation are shown. GlcNAc residues may be N-deacetylated and N-sulfated. In chondroitin sulfate and heparan sulfate, the extent of these modifications is highly variable, and the combination of variably modified disaccharide units leads to the huge structural diversity of these glycosaminoglycans

4.2.2 The Binding of Pleiotrophin/Midkine with Phosphacan It has been demonstrated that various cell adhesion molecules (Nr-CAM, L1/Ng-CAM, F3/contactin, N-CAM, and TAG-1), heparin-binding growth factors (FGF-2, pleiotrophin, and midkine), and ECM molecules (tenascin-C and tenascin-R) bind with the extracellular region of PTPz (Maeda, 2007). However, only pleiotrophin and midkine have been shown to function as signal-transducing ligands for PTPz. Pleiotrophin and midkine are composed of two domains that are held by disulfide bridges and are rich in basic amino acids (Muramatsu, 2002). These proteins share 45% amino acid sequence identity, promote survival, neurite outgrowth, and migration of neurons, and play important roles in tissue remodeling and carcinogenesis (Muramatsu, 2002). Pleiotrophin and midkine show similar low- (Kd = 3 nM) and high- (Kd = 0.2  0.6 nM) affinity bindings to phosphacan (Maeda et al., 1996, 1999). Removal of chondroitin sulfate by chondroitinase ABCtreatment decreases the binding affinity of phosphacan to both pleiotrophin and midkine. In contrast to the intact phosphacan, the chondroitinase-treated molecule shows a single low-affinity binding to these growth factors (Kd = 13 nM). This suggests that the chondroitin sulfate portion of phosphacan plays critical roles in determination of the binding affinity for pleiotrophin and midkine, and that there are at least two

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subpopulations of phosphacan bearing chondroitin sulfate chains with different structures. Our recent studies suggested that the density of D- (GlcA(2S)b1–3GalNAc(6S)) and E-type (GlcAb1–3GalNAc (4S, 6S)) oversulfated structures in chondroitin sulfate determine the affinity of phosphacan for pleiotrophin (> Figure 11-5). That is, phosphacan with oversulfated structure-rich chondroitin sulfate displays

. Figure 11-5 Schematic model of PTPz signaling. Pleiotrophin binds to the extracellular domain of PTPz, in which highaffinity binding requires oversulfated structures in chondroitin sulfate (D or E type structures). Pleiotrophin binding leads to the dimerization or oligomerization of PTPz, resulting in the inactivation of PTP activity. Phosphacan may promote the signaling as a reservoir of pleiotrophin or inhibit it as a competitive inhibitor of ligand binding, depending on the structure of chondroitin sulfate

higher affinity for pleiotrophin than that without these structures (Maeda et al., 2003, 2006). The structure of the chondroitin sulfate on phosphacan dynamically changes during development of the brain, suggesting that this structural change regulates the signaling of pleiotrophin and midkine (Maeda et al., 2003; Shimazaki et al., 2005).

4.2.3 Pleiotrophin Is a Ligand of PTPz It has been reported that pleiotrophin-PTPz signaling is involved in the neurite extension of cortical neurons, the morphogenesis of cerebellar Purkinje cell dendrites, and the cell migration of cortical neurons, glioblastoma cells, and endothelial cells (Maeda et al., 1996; Maeda and Noda, 1998; Muller et al., 2003; Tanaka et al., 2003; Lu et al., 2005; Polykratis et al., 2005; Ulbricht et al., 2006). Midkine-PTPz signaling has

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also been shown to regulate the survival of embryonic cerebral neurons and the cell migration of cortical neurons and osteoblast-type cells (Maeda et al., 1999; Qi et al., 2001; Sakaguchi et al., 2003). Many of these signaling events were inhibited by the addition of chondroitin sulfate preparations (Maeda and Noda, 1998; Maeda et al., 1999; Qi et al., 2001; Tanaka et al., 2003; Polykratis, 2005) and chondroitinase ABC digestion (Qi et al., 2001; Sakaguchi et al., 2003; Tanaka et al., 2003), indicating that the chondroitin sulfate portion of PTPz is essential for the signaling. It is notable that the effects of chondroitin sulfate preparations are structure dependent. Pleiotrophin/midkine-PTPz signaling was strongly inhibited by D-type structure-rich chondroitin sulfate C and D and by E-type structure-rich chondroitin sulfate E, but not by chondroitin sulfate A, which contains almost no oversulfated structures (Maeda and Noda, 1998; Maeda et al., 1999; Qi et al., 2001; Tanaka et al., 2003; Polykratis et al., 2005). This is consistent with the finding that chondroitin sulfate C, D, and E but not chondroitin sulfate A inhibit the binding between phosphacan and pleiotrophin/ midkine (Maeda et al., 1996, 1999). In the developing brain, the content of phosphacan is much higher than that of PTPz (Nishiwaki et al., 1998). At present, the functional relationship between PTPz and phosphacan is unclear. However, the finding that oversulfated structures in the chondroitin sulfate on phosphacan regulate its binding affinity for pleiotrophin suggests one possibility (> Figure 11-5). Phosphacan bearing highly oversulfated chondroitin sulfate might competitively inhibit the binding of pleiotrophin to PTPz, and therefore suppress the signaling. On the other hand, phosphacan with a low content of oversulfated structures might support the signaling by serving as a reservoir of pleiotrophin.

4.2.4 Pleiotrophin Induces Oligomerization of PTPz Several reports demonstrated that pleiotrophin stimulation increases tyrosine phosphorylation of substrate molecules of PTPz by suppressing its PTP activity (Meng et al., 2000; Kawachi et al., 2001; Pariser et al., 2005). Sodium channels, Cat-1/GIT1, b-catenin, and b-adducin were identified as substrates of PTP z , and the tyrosine phosphorylation of the latter three proteins was shown to be increased after pleiotrophin stimulation (Meng et al., 2000; Ratcliffe et al., 2000; Kawachi et al., 2001; Pariser et al., 2005). A recent study showed that pleiotrophin promotes oligomerization of PTPz on the cell surface, which leads to the inactivation of its PTP activity (Fukada et al., 2006), indicating that the signal transduction of PTPz proceeds according to the dimerization-based model (> Figure 11-2b). The functional role of the chondroitin sulfate portion of PTPz in the oligomerization process is unclear. However, such a process should be highly dependent on the chondroitin sulfate structure including the chain length and sulfation pattern. In the presence of a large amount of high-affinity phosphacan, which works as a competitive inhibitor, oversulfated chondroitin sulfate on PTPz must be necessary for pleiotrophin binding and oligomerization (high-affinity receptor form) (> Figure 11-5). However, when the concentration of phosphacan is low or the pleiotrophin concentration is high, oversulfated chondroitin sulfate on PTPz may not be needed (low-affinity receptor form). In addition, the chain length of chondroitin sulfate might have an important regulatory function, because long chondroitin sulfate chains on PTPz could sterically hinder the process of oligomerization of the receptor. Thus, it seems likely that multiple factors, including the sulfation pattern and chain length of chondroitin sulfate on PTPz and phosphacan and the balance of the concentrations of PTPz, phosphacan, and pleiotrophin, determine the strength of signal transduction (> Figure 11-5). Such complex regulation of the receptor function should contribute to the fine-tuning of delicate cell–cell interactions in the developing nervous system.

4.3 DLAR and Neural Functions 4.3.1 Drosophila RPTPs and Glycosaminoglycans The Drosophila genome encodes six RPTP genes: two type IIA RPTPs (DLAR and DPTP69D) and four type III RPTPs (DPTP99A, DPTP10D, DPTP52F and DPTP4E). While DPTP4E is widely expressed, the others

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are selectively expressed in the nervous system ( Johnson and Van Vactor, 2003). As described earlier, there are 21 vertebrate RPTP genes, which are classified into 8 types. Thus, the Drosophila genome shows a very simple composition of RPTP genes, which may reflect the simple structure of the Drosophila body compared with those of vertebrates. Similarly, glycosaminoglycans in Drosophila show simple structures compared with those of vertebrates. Although heparan sulfate in Drosophila exhibits structural heterogeneity comparable with that of vertebrates, the structure of Drosophila chondroitin sulfate is extremely simple (Toyoda et al., 2000). Chondroitin sulfate in Drosophila is extremely undersulfated and contains only O (GlcAb1–3GalNAc) and A type (GlcAb1–3GalNAc(4S)) structures. On the other hand, vertebrate chondroitin sulfate contains C (GlcAb1– 3GalNAc(6S), D, and E types and other structures in addition to the O and A type structures, as described earlier. Thus, although it seems that heparan sulfate plays similar roles in both Drosophila and vertebrates, such as regulation of growth factor signaling, Drosophila chondroitin sulfate is not likely to be involved in signaling function. So, it may be that vertebrates diversified RPTP and glycosaminoglycan structures to accomplish diverse cell–cell and cell–matrix interactions that are necessary for the development of their complex body structures.

4.3.2 RPTP and Motor Axon Guidance in Drosophila In Drosophila embryos, motor axons exit laterally from the central nervous system and navigate to their muscle targets along five nerve fascicles in a stereotyped manner (ISN, ISNb, SNa, SNc, and SNd). These motor axons appear to express all or most of the neural RPTPs (DLAR, DPTP69D, DPTP99A, DPTP10D, and DPTP52F) (Schindelholz et al., 2001). A series of experiments analyzing motor axon guidance in single or multiple RPTP mutants indicated that a specific combination of neural RPTPs are required for each axon to correctly navigate to the target (Desai et al., 1996, 1997; Krueger et al., 1996; Schindelholz et al., 2001). These RPTPs competitively or cooperatively work in the growth cones to select the correct pathways, and the relationships among RPTPs seem to be dependent on the cellular context. Namely, the same combination of RPTPs can act either cooperatively or competitively at growth cone choice points depending on the motor nerves and their developmental stages (Schindelholz et al., 2001). It is considered that specific ligands for particular RPTPs are localized at each growth cone choice point and the RPTPs are activated or inactivated by these ligands leading to the axon guidance decision (Fox and Zinn, 2005).

4.3.3 DLAR and Syndecan in Motor Axon Guidance In Drosophila embryos, ISNb motor axons leave the common ISN pathway at the ‘‘exit junction,’’ enter the ventral target region, and then form synapses on specific ventral muscles. On the other hand, in DLAR mutant embryos, ISNb axons fail to enter the ventral target region and continue to grow in a bundle adjacent to the common ISN pathway (Krueger et al., 1996; Fox and Zinn, 2005). The extracellular region of DLAR is composed of three immunoglobulin-like domains and nine fibronectin type III repeats, among which the first immunoglobulin-like domain contains basic amino acid clusters that constitute the heparinbinding site (Fox and Zinn, 2005). Biochemical studies indicated that DLAR binds with the heparan sulfate chains of syndecan through the heparin-binding site with subnanomolar affinity, and genetic interaction studies using loss-of-function mutants of syndecan and DLAR further demonstrated that syndecan contributes to DLAR function during ISNb axon guidance (Fox and Zinn, 2005). Further studies revealed that syndecan in muscle serves as a ligand of DLAR on SNa motor axons (Fox and Zinn, 2005).

4.3.4 Interaction of DLAR with Syndecan and Dallylike By the time of larval hatching, Drosphila motor nerves form nascent neuromuscular junctions in the periphery, and then extend multiple branches decorated with many synaptic boutons that contain active

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zones during larval life ( Johnson et al., 2006). In addition to axon guidance in Drosophila embryos, it has been demonstrated that DLAR is required to develop normal synaptic morphology and to define the shape and size of the active zone in larvae (Kaufmann et al., 2002). Johnson et al. (2006) showed that Drosophila glypican Dallylike as well as syndecan binds with DLAR. The heparan sulfate portion of Dallylike binds to the immunoglobulin-like domains of DLAR, suggesting that Dallylike and syndecan interact with a common heparin-binding site in the first immunoglobulin domain of DLAR (Fox and Zinn, 2005; Johnson et al., 2006). Furthermore, Scatchard analyses indicated that syndecan and Dallylike competitively bind with DLAR with similar nanomolar affinities (Kd = 13 nM for syndecan and Kd = 8 nM for Dallylike) (Johnson et al., 2006). The localization of both syndecan and Dallylike are specific to the synapses, whereas motor axons and bouton-free regions of presynaptic arbors show little or no expression, suggesting that interactions between DLAR and these heparan sulfate proteoglycans play roles in the synaptogenesis.

4.3.5 DLAR, Syndecan, and Dallylike in Synaptogenesis Although both syndecan and Dallylike bind to DLAR with similar affinity, their functions are quite different (Johnson et al., 2006). The function of syndecan is limited to the growth of synapses and is not related to morphogenesis of the active zone, whereas Dallylike contributes to the active zone morphogenesis but not to synapse growth. Genetic analysis indicated that syndecan and DLAR act in a common signaling pathway that regulates the synaptogenesis. It seems that syndecan associates with DLAR at the presynaptic membrane of motor axons and activates cell-autonomously the PTP activity of DLAR, which leads to growth of synaptic boutons (> Figure 11-6). On the other hand, Dallylike appears to inactivate the PTP activity of . Figure 11-6 DLAR signaling. Presynaptic syndecan in association with DLAR promotes the growth of synaptic boutons (left). Dallylike inactivates DLAR, increases tyrosine phosphorylation of enabled (Ena) and stabilizes active zones (right). Syndecan and Dallylike competitively bind to DLAR immunoglobulin-like domain and mediate a transition between synapse growth and active zone assembly

Receptor-like protein tyrosine phosphatases and proteoglycans in the nervous system

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DLAR, which leads to the active zone stabilization (> Figure 11-6) (Johnson et al., 2006). Further in vitro experiments indicated that Dallylike inhibits DLAR activity and increases the tyrosine phosphorylation level of Enabled (Ena), a substrate of DLAR (> Figure 11-6) (Johnson et al., 2006). From these findings, it has been proposed that presynaptic growth is initially promoted by syndecanDLAR signaling and then halted by competitive binding of Dallylike to DLAR, which results in functional maturation of synapses (> Figure 11-6) (Johnson et al., 2006). The affinities of many heparin-binding proteins for heparan sulfate are dependent on the structure of heparan sulfate (Bandtlow and Zimmermann, 2000). Thus, competitive binding of syndecan and Dallylike to DLAR may be regulated by the differential modification of heparan sulfate on these proteoglycans. Dunah et al. (2005) demonstrated that vertebrate LAR also plays important roles in development and maintenance of hippocampal excitatory synapses. It will be interesting to know whether syndecan and glypican are involved in LAR signaling.

5

Conclusion

Recent studies demonstrated that RPTPs and proteoglycans play important roles in the development and plasticity of the nervous system. However, many problems remain to be solved. Especially, ligands and substrate molecules of RPTPs are still unknown for many RPTPs. Furthermore, little is known about the structure-dependent function of glycosaminoglycans in the nervous system. Recent investigations indicated that chondroitin sulfate proteoglycans and heparan sulfate proteoglycans have somewhat overlapping functions. Namely, chondroitin sulfate can associate with heparin-binding sites of some proteins that have been regarded as only heparan sulfate-binding proteins (Maeda et al., 1996, 1999). The extracellular regions of several RPTPs bind with heparin (Aricescu et al., 2002; Fox and Zinn, 2005), so competitive binding of heparan and chondroitin sulfate proteoglycans to RPTPs may play important regulatory functions. Future studies should reveal the unexpected functions of these diverse molecules in the nervous system.

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The Metabolism and the Functions of Diphosphoinositol Polyphosphates

S. B. Shears

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 226

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Discovery and Nomenclature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 226

3 The Enzymology of Diphosphoinositol Polyphosphate Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229 3.1 The Kinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229 3.2 The Phosphatases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231 4 Proposed Mechanisms of Action of Diphosphoinositol Polyphosphates . . . . . . . . . . . . . . . . . . . . . . . . 233 4.1 A Novel Mode of Posttranslational Modification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 4.2 Competitive Ligand Binding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 234 5 5.1 5.2 5.3 5.4 5.5

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Biological Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 DNA Maintenance and Repair . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 235 Yeast Vacuole Biogenesis and Vesicle Trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 236 Apoptosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 236 Transcriptional Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237 Exocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238

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The metabolism and the functions of diphosphoinositol polyphosphates

Abstract: This review describes current understanding of the metabolism and the functions of a subgroup of the inositol phosphate signaling family which contain diphosphate groups (the diphosphoinositol polyphosphates, often called “inositol pyrophosphates”). The review begins with a short historical perspective of this field of research. There is a section on the enzymology of the kinases and phosphatases that are responsible for regulating the turnover of these polyphosphates. There is a discussion of the current status of our rather limited insight into the mechanisms of action of the diphosphoinositol polyphosphates. Finally, there is a review of several of the proposed biological functions of these polyphosphates. List of Abbreviations: CK2, casein kinase 2; CRAC, cytosolic regulator of adenylyl cyclase; DIPPs, diphosphoinositol polyphosphate phosphohydrolases; ENTH, epsin N-terminal homology; PIKKs, phosphoinositide 3-kinase related kinases

1

Introduction

The phosphorylated inositol moiety has been described as a fundamental signaling entity that the cell utilizes to generate combinatorially complex arrays of communication pathways with multiple functions (York and Hunter, 2004). These molecules exist as entities that are either soluble (as inositol phosphates) or membrane bound (the inositol lipids, or phosphoinositides). Yet, for reasons that are still being debated (Irvine and Schell, 2001; Shears, 2007), the inositol lipids have, for many years, far outpaced the inositol phosphates in the ratings war. In fact, it is not even that unusual for the inositol phosphates to be falsely accused of merely being another “phosphoinositide” (e.g., Liu et al., 2004). This certainly obscures the fact that the physicochemical properties and biological actions of the two groups of signals are very different. Fortunately—not in the least for those of us who have invested their research careers in this topic—the last few years have seen a number of high-profile publications in the inositol phosphate field, finally signaling the long-awaited (Irvine and Schell, 2001) revival of interest in this functionally wide-ranging group of cellular messengers. This renaissance should be of particular benefit to research into one specialized subgroup of inositol phosphates: the diphosphoinositol polyphosphates. Up to 2005, 15 years after the discovery of these molecules (Menniti et al., 1993; Stephens et al., 1993), less than five papers a year were devoted to this topic (Shears, 2007). This situation led one inositide expert to note that research in this area “. . .has gone somewhat cold” (Berridge, 2006), a statement which rather obscured the fact that it had not yet been that hot. But, change is in the air. The diphosphoinositol polyphosphates are being carried along by the new wave of interest in inositol phosphates as important signaling entities. In particular, there has been considerable general interest in demonstrations that diphosphoinositol polyphosphates engage in a novel form of covalent modification, namely, the kinase-independent diphosphorylation of proteins (Saiardi et al., 2004; Bhandari et al., 2007). We shall look closely at this concept in this chapter. Furthermore, the molecular identification of the kinases that synthesize the diphosphoinositol polyphosphates (Saiardi et al., 1999, 2001; Schell et al., 1999; Luo et al., 2003; Choi et al., 2007; Fridy et al., 2007; Mulugu et al., 2007) has engendered a number of genetic studies into the biological functions of these polyphosphates. As a result, a number of physiological processes are now considered targets for regulation by diphosphoinositol polyphosphates. These include vesicle trafficking (Saiardi et al., 2002), transcription (El Alami et al., 2003; Lee et al., 2007), chemotaxis (Luo et al., 2003), telomere maintenance (Saiardi et al., 2005; York et al., 2005), apoptosis (Morrison et al., 2001; Nagata et al., 2005), environmental stress responses (Dubois et al., 2002; Pesesse et al., 2004; Safrany, 2004; Choi et al., 2005), and DNA repair (Luo et al., 2002). These reports will also be discussed here.

2

Discovery and Nomenclature

As the 1980s drew to a close, it seemed that there were two separate classes of inositol phosphates (Berridge and Irvine, 1989). One, the so-called “intracellular signaling” group, included Ins(1,4,5)P3 and Ins(1,3,4,5)P4; these are present at low levels in resting cells (0.1–1 mM), but they rapidly increase in concentration following receptor activation (Berridge and Irvine, 1989). The other group, defined as “agonist

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insensitive,” included Ins(1,3,4,5,6)P5 and InsP6, which are clearly present at higher levels (around 50 mM), but did not initially appear to respond to short-term receptor activation (Berridge and Irvine, 1989; French et al., 1991). The metabolism of InsP6 in particular was, at that time, thought to be especially lethargic. A perfectly logical reason underpinned this interpretation: it had been noted that, in mammalian cells, it takes several days radiolabeling with [3H]inositol before the cellular pool of [3H]InsP6 attains equilibrium labeling (Michell et al., 1990; French et al., 1991). Thus, InsP6 was initially not thought to be of any significance to signal transduction (Berridge and Irvine, 1989). Instead (other than when it had attracted somewhat fleeting interest as an antioxidant (Berridge and Irvine, 1989)), InsP6 was suggested to serve mainly as a phosphate store, an idea that was borrowed from the plant literature (Michell, 1986). By the early 1990s, several groups had noted that mammalian cells contained low levels of inositol derivatives that are more polar than InsP6 (Stephens et al., 1991; Oliver et al., 1992; Wong et al., 1992). Then came the startling observation that Dictyostelium synthesized these novel molecules at levels that were almost as high as InsP6 itself (Europe-Finner et al., 1991). Clearly, inositol phosphate research was set to enter a new phase. These preliminary observations culminated in 1993 in the formal identification of these new metabolites as diphosphorylated derivatives of Ins(1,3,4,5,6)P5 and InsP6 (Menniti et al., 1993; Stephens et al., 1993) (see > Figure 12-1). We (Menniti et al., 1993) originally classified these molecules as “inositol pyrophosphate polyphosphates,” but it became common practice to abbreviate this too, simply as, “inositol pyrophosphates” (e.g., Saiardi et al., 2002). IUPAC frowns on this vernacular terminology (IUPAC-IUB Commission on Biochemical Nomenclature, 1977). It is understandable that there might be some indifference to IUPAC’s disapproval, since the politically correct nomenclature—“diphosphoinositol polyphosphates” (Stephens et al., 1993)—does not exactly roll off the tongue. However, to be fair, “inositol pyrophosphate” is a designation that does not adequately describe a molecule that contains both phosphates and diphosphates, nor does it embrace triphosphosphates, which is also a genuine concern (see later). Hence, the long-winded terminology is used in this chapter. The reader should also note that the diphosphorylated derivatives of InsP6, namely, PP-InsP5 and [PP]2-InsP4, are frequently abbreviated in the literature as “IP7” and “IP8,” respectively (e.g., see > Figure 12-1). This terminology is avoided here, for fear of diverting attention away from the diphosphate derivatives of Ins(1,3,4,5,6)P5, which may have their own, separate functions (> Section 5.1). With the notable exception of Dictyostelium (now considered to be awash with 100–250 mM concentrations of diphosphoinositol polyphosphates (Laussmann et al., 2000)), most current estimates put the levels of these molecules in the 0.5–5 mM range in yeast and mammalian cells (Fisher et al., 2002; Ingram et al., 2003; Barker et al., 2004; Bennett et al., 2006; Illies et al., 2007). This may not sound like much, but that concentration range is similar to that seen for other inositol phosphate signals such as Ins(1,4,5)P3 and Ins(1,3,4,5)P4. The low levels of diphosphoinositol polyphosphates in cells makes their analysis a little laborious. There is a nonradioactive, automated, in-line HPLC assay for inositol phosphates that relies on a postcolumn formation of a metal–dye complex, but its sensitivity is not adequate for studying [PP]2-InsP4 turnover in mammalian cells (Albert et al., 1997). Instead, most laboratories use an [3H]inositol-labeling procedure. The diphosphoinositol polyphosphates can only be detected after cells are radiolabeled with [3H]inositol for several days, because of the slow accumulation of [3H]inositol by the Ins(1,3,4,5,6)P5 and InsP6 precursors. This makes their analysis expensive. It is also a laborious undertaking, because HPLC is required in order to separate the individual [3H]-labeled inositol phosphates. Even worse, in-line scintillation counters lack the sensitivity necessary to accurately assay [3H]-labeled diphosphoinositol polyphosphates in the high-salt buffers that are required to elute them from the HPLC column. It is essential to use a fraction collector, manually add scintillant to each fraction, mix vigorously (Azevedo and Saiardi, 2006), and use a traditional counter. Anyone who doubts the veracity of the latter statement should note “the lack of observable in vivo kinase activity,” when an in-line counter failed to detect increases in [3H]-labeled diphosphoinositol polyphosphates following overexpression of the PP-InsP5 kinase in HEK cells (Fridy et al., 2007). In contrast, using the same cell line, the collection and analysis of individual fractions clearly identifies a small [PP]2-InsP4 peak in nontransfected cells (0.2% of the InsP6 peak), which was elevated 30-fold following overexpression of the PP-InsP5 kinase (Choi et al., 2007).

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The metabolism and the functions of diphosphoinositol polyphosphates

. Figure 12-1 The diphosphoinositol polyphosphates: nomenclature and diversity. In the abbreviations of the chemical structures, “Ins” indicates the myo-inositol skeleton. The number of monophosphates around the inositol ring is denoted as a suffix after the “P.” The prefixes denote the number of diphosphates (PP). The schematic shows reactions catalyzed by the InsP5 kinase (purple arrow; also known as IPK1), the InsP6 kinases (“IP6K”; green arrows), and the PP-InsP5 kinases (“PPIP5K” or “VIP”; blue arrows). One of the PP-InsP5 isomers has the diphosphate group attached to the 5-carbon (Albert et al., 1997). The other PP-InsP5 isomer is now (Lin, Fridy, Choi, Shears, York and Mayr, unpublished data) believed to have the diphosphate attached to the 1 or 3 position (3-PP-InsP5 is shown in the figure). Mammalian [PP]2-InsP4 has diphosphates at the 5- and 1/3-positions (Lin, Fridy, Choi, Shears, York and Mayr, unpublished data). In Dictyostelium it has been reported that 5,6[PP]2-InsP4 (Laussmann et al., 1997), and 1/3,5-[PP]2-InsP4 are both present (Laussmann et al., 1998). All InsP6 kinases also phosphorylate InsP5 to PP-InsP4 (Saiardi et al., 2000). PP-InsP4 is itself further phosphorylated by the InsP6 kinase, but the product is unclear. It was originally proposed to be [PP]2-InsP3 (Saiardi et al., 2000), but [PPP]-InsP4 is a possible alternate product (see (Shears, 2004)). Similarly, the further phosphorylation of PP-InsP5 by the “InsP6 kinase” might yield [PPP]-InsP5 rather than the [PP]2-InsP4 that was initially presumed (Saiardi et al., 2000)

The diphosphoinositol polyphosphates turn over rapidly (Menniti et al., 1993). The latter observation arose from the addition of fluoride to intact cells (Menniti et al., 1993). This inhibits the diphosphoinositol polyphosphate phosphohydrolases (DIPPs) (Safrany et al., 1998), exposing the otherwise well-disguised, ongoing activities of the InsP6 and PP-InsP5 kinases (> Figure 12-1). This causes cellular levels of PP-InsP4, PP-InsP5, and [PP]2-InsP4 to increase substantially (the reader who is tempted to refer to our 1993 study (Menniti et al., 1993) will note that the structure of [PP]2-InsP4 was not then appreciated, and so it was labeled as “IP6X,” to depict the “unknown derivative of InsP6”). This fluoride-dependent increase in levels of diphosphoinositol polyphosphates is accompanied by substantial decreases in the levels of the Ins(1,3,4,5,6)P5 and InsP6 precursors (Menniti et al., 1993; Safrany and Shears, 1998). This demonstration of the high metabolic activity of the diphosphoinositol polyphosphates has been one of the driving forces

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behind research in this field; Safrany et al. (1999) have argued the cell must recoup some benefit from the substantial investment of cellular energy needed to maintain these active substrate cycles. Another notable feature of this class of polyphosphates is that the hydrolysis of their diphosphate groups has been viewed as a “high-energy” reaction that ought to have particular bioenergetic significance (Stephens et al., 1993; Laussmann et al., 1996; Voglmaier et al., 1996). The latter idea now takes much of the responsibility for the current upsurge in general interest in this field (> Section 4.1). Nevertheless, the roles of Ins(1,3,4,5,6)P5 and InsP6 should not be viewed only in terms of their being precursor pools for the diphosphoinositol polyphosphates. In particular, InsP6 has attracted its own band of enthusiasts from within the cell-signaling community. For example, it became clear that if the cellular pool of InsP6 was equilibrium labeled with [3H]inositol, substantial stimulus-dependent changes in its levels could then be observed, albeit typically rather long term, such as those that occur during cell differentiation (French et al., 1991). A recent report indicating the existence of hidden, and metabolically superactive subpools of InsP6 (Otto et al., 2007) suggests that there is still much more for us to learn about this polyphosphate. Many intracellular roles have been attributed to InsP6 (for reviews, see Irvine and Schell, 2001; Shears, 2001; York, 2006; Alcazar-Roman and Wente, 2008). However, that is a subject beyond the remit of this chapter.

3

The Enzymology of Diphosphoinositol Polyphosphate Metabolism

3.1 The Kinases In mammalian cells, there are three isoforms of kinases (types 1, 2, and 3) that phosphorylate both Ins (1,3,4,5,6)P5 and InsP6, and they have a molecular weight of 46–49 kDa (Saiardi et al., 1999, 2001; Schell et al., 1999). These kinases possess a PxxxDxKxG catalytic domain, and a remote, catalytically essential SSLL (or similar) tetrapeptide. Some homologues of these kinases are rather larger, such as the 120-kDa protein in yeast (Saiardi et al., 1999) and the 72-kDa kinase in Dictyostelium (Luo et al., 2003). The kinase in S. cerevisiae (Kcs1) is unique in possessing two leucine zipper motifs. These are amphipathic heptad repeats, (LxxxxxxL)n, where the value of “n” is typically 4 or 5, which form coiled-coil structures that dimerize; leucine zippers are found in proteins that regulate transcription (Vinson et al., 2002). Indeed, there is evidence that the yeast InsP6 kinase activity does influence gene expression (> Section 5.4). These enzymes are near-universally known as “InsP6 kinases,” often abbreviated to “IP6K,” despite their physiologically relevant ability to also actively phosphorylate Ins(1,3,4,5,6)P5. Acknowledging this fact, we (Dubois et al., 2002) and others (York et al., 2005; Alcazar-Roman and Wente, 2008) have proposed that “diphosphoinositol polyphosphate synthase” is a more appropriate nomenclature. However, this proposal has not been widely adopted. A demonstration that purified, recombinant InsP6 kinase can further phosphorylate PPInsP5, in vitro at least (Saiardi et al., 2000), has inspired speculation (Shears, 2004; Shears, 2005) that the product might conceivably be a triphosphate, PPP-InsP5, rather than the bis-diphosphoinositol, [PP]2InsP4 (> Figure 12-1). It might still be profitable to explore this possibility. It has been shown that the IP6Ks selectively synthesize a diphosphate group at the 5-phosphate of InsP6 (Mulugu et al., 2007); in mammals at least, 5-PP-InsP5 is also the predominant isomer to accumulate inside cells (Albert et al., 1997). It is presumed—but has yet to be formally proven—that Ins(1,3,4,5,6)P5 is also phosphorylated at the 5-position by the InsP6 kinase. Most studies agree that these kinases display a Km for InsP6 in the range of 0.4–2 mM, with an approximately tenfold lower affinity for Ins(1,3,4,5,6)P5 (Menniti et al., 1993; Voglmaier et al., 1996; Saiardi et al., 2000, 2001). The fact that IP6K activity can be regulated has been most dramatically demonstrated with Dictyostelium: when the free-living amoeboid form of this organism exhausts its food supply and aggregates into a multicellular organism, the cellular levels of PP-InsP5 increase tenfold within a few hours (Laussmann et al., 2000). This effect occurs through the actions of a G-protein-coupled cAMP receptor (Luo et al., 2003). In mammalian cells, release of [Ca2+] from the endoplasmic reticulum slightly inhibits cellular InsP6 kinase activity (Glennon and Shears, 1993), but neither the underlying mechanisms, nor the biological significance of this effect of Ca2+, have been ascertained.

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This area of research took a new turn when, in 2005, York et al. noted an apparently upregulated InsP6 kinase activity in a strain of S. cerevisiae from which the genes encoding Kcs1 and the diphosphoinositol polyphosphate phosphatase (Ddp1) had both been eliminated. The purification and cloning of this InsP6 kinase activity in an elegant biochemical study revealed a protein (named Vip1) that is very different from the canonical IP6Ks (Mulugu et al., 2007). This new kinase is a 120-kDa protein with three roughly equally sized sections. The N-terminal region contains the kinase domain (Mulugu et al., 2007). The middle section represents a putative “histidine-acid phosphatase” domain. However, this is a somewhat puzzling observation because no actual phosphatase activity has been observed yet. The function of the remaining C-terminal region of the protein is unknown at this time. Why was this kinase named Vip1? This originates, rather tangentially, from an earlier study of the homologue in Schizosaccharomyces pombe, Asp1, prior to it being realized that this protein is an inositol phosphate kinase. Instead, the name originates from the discovery of genetic interactions between Asp1 and genes encoding several cytoskeletal proteins: Arp, Sop, and Profilin (Feoktistova et al., 1999). However, the homologue in S. cerevisiae could not be awarded the same name, because that yeast already possessed an Asp1 gene (encoding L-asparaginase). Since an asp is also a snake of the Viperidae family, Vip1 became the logical second choice for the christening of the S. cerevisiae homologue (Gould, personal communication). We have independently purified, sequenced, and cloned two mammalian PP-InsP5 kinases that turned out to be homologues of Vip1 (Choi et al., 2007). At the same time, York’s group separately recognized the nature of the mammalian genes by homology-based cloning (Fridy et al., 2007). Both in vivo and in vitro, the PP-InsP5 kinase activities of the mammalian enzymes appeared to be quantitatively more important than the InsP6 kinase activity (Choi et al., 2007). As a result, there now seems little doubt that these are the PP-InsP5 kinases that we and others have been trying to clone for many years (Shears et al., 1995; Huang et al., 1998). As IUPAC had already decided that “diphosphoinositol pentakisphosphate kinase” (E.C. 2.7.4.24) is an appropriate name for these enzymes, my laboratory introduced PPIP5K1 and PPIP5K2 as their abbreviations (Choi et al., 2007). This nomenclature is also consistent with InsP6 kinase frequently being abbreviated to “IP6K.” A 1-D NMR analysis revealed that the isomer of PP-InsP5 that is synthesized by Vip1 is different from the 5-PP-InsP5 made by the canonical IP6Ks (Mulugu et al., 2007). The authors (Mulugu et al., 2007; York and Lew, 2008) did take care to point out that the nature of their structural analysis did not allow them to determine whether the diphosphate on this new isomer of PP-InsP5 was added to either the 1/3 or the 4/6 positions (the alternatives within each pair are enantiomers that are also technically difficult to distinguish between). However, more recent studies (Lin, Fridy, Choi, Shears, York and Mayr, unpublished data) have pursued this problem and determined that both yeast Vip1 and its human homologues add a diphosphate group to the 1/3 position. Thus, the [PP]2-InsP4 in mammalian and yeast cells is now known to have diphosphate groups at both the 1/3 and the 5-positions. The Vip1/PPIP5K enzymes can also phosphorylate InsP6 to 1/3-PP-InsP5. The latter’s further phosphorylation by the 5-kinase activity of the IP6Ks provides a second route by which 1/3,5-[PP]2-InsP4 can be synthesized (Mulugu et al., 2007). The same isomer of [PP]2-InsP4 can also be synthesized by the Dictyostelids (Laussmann et al., 1998). Interestingly, Dictyostelium also contain 6-PP-InsP5 and 5,6-[PP]2-InsP4 (Laussmann et al., 1997). While the exact function of [PP]2-InsP4 remains to be determined, it is already known that the 1/3-PP-InsP5 synthesized by the InsP6 kinase activity of Vip1 is biologically important. For example, a BLAST search of the Arabidopsis genome (October 2007) failed to uncover any homologues of IP6K, but a candidate Ppip5k/Vip1 homologue is present; the latter enzyme may offer the only route by which Arabidopsis can synthesize PP-InsP5. Moreover, the novel isomer of PP-InsP5 synthesized by the InsP6 kinase activity of Ppip5k/Vip1 may have an important function in yeast (> Section 5.4). The PPIP5Ks/VIPs display a Km value of 0.1–0.5 mM for PP-InsP5 as substrate (Choi et al., 2007; Fridy et al., 2007). Thus, in mammalian cells, the enzymes are likely operating at less-than-maximal capacity, since the cellular levels of PP-InsP5 are generally within the 0.6–5 mM range (Fisher et al., 2002; Ingram et al., 2003; Barker et al., 2004; Illies et al., 2007), with the notable exception of the Dictyostelids (> Section 3.1). Against a background of there being substantial interest in understanding why all of the other inositol phosphates kinases can be found in the nucleus (Alcazar-Roman and Wente, 2008; Seeds and York, 2007), it was, perhaps, surprising to find that the mammalian PPIP5Ks are entirely cytoplasmic (Choi et al., 2007; Fridy et al., 2007).

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The cloning of the PPIP5Ks that synthesize [PP]2-InsP4 has connected what was otherwise an important missing link in this field of research (Shears, 2007). Moreover, the PPIP5Ks are regulated enzymes, and their molecular characterization (Choi et al., 2007; Fridy et al., 2007; Mulugu et al., 2007) will now help us pursue the mechanisms involved. For example, cAMP inhibits [PP]2-InsP4 synthesis by a novel, but unexplained mechanism that does not involve protein kinases A or G (Safrany and Shears, 1998). Moreover, we have previously shown that the levels of [PP]2-InsP4 increase up to 20-fold when cells are exposed to hyperosmotic stress (Pesesse et al., 2004; Choi et al., 2007). [PP]2-InsP4 levels also increase during a thermal challenge (either heating or cooling), although much less dramatically (Choi et al., 2005). Finally, in S. cerevisiae, the kinase activity of Ppip5K/Vip1 toward InsP6 was recently reported (Lee et al., 2007) to increase significantly when yeast are transferred to a low-phosphate medium; within a couple of hours, cellular levels of PP-InsP5 increased tenfold (but see > Section 5.4). Even though PPIP5Ks are only known to be acutely regulated in just these relatively limited set of conditions, the data nevertheless indicate that these enzymes are important in cell signaling and, therefore, deserve further study. Understanding the role of PPIP5Ks during hyperosmotic stress is a particular focus of my laboratory. One problem with studying this subject is that it has been a popular opinion for much of the previous 70 years (Darrow and Yannet, 1935; Bourque and Oliet, 1997) that metazoan cells, with the exception of those of the renal medulla, are largely protected from the potentially deleterious effects of anisosmotic gradients, by virtue of their being bathed in an osmotically stable extracellular fluid (280–300 milliosmol/kg in most mammals). However, this long-standing consensus is now being successfully challenged by more recent research: several nonrenal cell types are now known to routinely experience changes in extracellular osmolarity, including airway epithelial cells, lymphocytes, and the various cell types in bones and cartilage (Knothe Tate, 2003; Go et al., 2004; Alfieri and Petronini, 2007). For example, lymphocyte development depends upon their ability to adapt to the hyperosmotic environment of the thymus (Go et al., 2004). Bones also represent an osmotically stressful environment: there are fluctuating osmotic pressure gradients between osteocytes and their surrounding proteoglycan-filled lacunocanalicular system which are important for mechanochemical coupling and for driving bulk fluid flow through the bone tissue (Knothe Tate, 2003). Hyperosmotic stress can also occur in airway epithelial cells when the composition of the airway surface liquid layer is compromised by inadequate airway humidification, such as during rapid breathing (e.g., during exercise), breathing of dry/cold air, tracheotomy breathing, and in some airway diseases (Song et al., 2001). Even in cell types that do not routinely experience these significant fluctuations in extracellular osmolarity, it is now accepted that mechanisms must be in place to adapt to the alterations in intracellular osmolarity that inevitably accompany normal cellular activities: ion transport across the plasma membrane, uptake and release of sugars and amino acids, and polymerization/depolymerization of macromolecules such as glycogen and proteins (Schliess and Haussinger, 2002). It is against this background that, hopefully, the reader will appreciate that it will be important to understand what is the significance of the cell allowing [PP]2-InsP4 levels to acutely increase following an osmotic shock (Pesesse et al., 2004; Choi et al., 2007). We originally proposed that the MEK/ERK pathway was responsible for stress-dependent activation of the PP-InsP5 kinase (Pesesse et al., 2004). In large part, this conclusion developed from the observation that the MEK inhibitors, U0126 and PD98059, both attenuated the stress-dependent elevations in cellular [PP]2-InsP4 levels (Pesesse et al., 2004; Choi et al., 2005). However, we subsequently determined that efficient “knockdown” of either MEK or ERK using RNA interference had not the slightest impact upon stress-dependent increases in [PP]2-InsP4 levels (Choi et al., 2008). Moreover, it now seems that the reason that U0126 and PD98059 inhibit cellular [PP]2-InsP4 synthesis is through an “off-target” perturbation of cellular adenine nucleotide balance (Dokladda et al., 2005), to which the PPIP5Ks are especially sensitive (Choi et al., 2008). Whether or not the latter effect itself has any biological significance remains to be determined.

3.2 The Phosphatases Cells contain active phosphohydrolases (DIPPs) that help ensure that there is rapid, ongoing turnover of diphosphoinositol polyphosphates. Five mammalian DIPPs have been cloned: type 1 (Safrany et al., 1998; Chu et al., 2004), types 2a/2b (Caffrey et al., 2000; Hua et al., 2001), and types 3a3b (Hidaka et al., 2002;

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Leslie et al., 2002; Hua et al., 2003). These five different DIPP enzymes do vary up to 25-fold in the values of their specificity constants (kcat/Km). Nevertheless, all of these values are exceptionally large, reaching 5
107 M1 s1 for PP-InsP5 hydrolysis by DIPP1, indicating that its activity is close to being limited by the diffusion-controlled encounter between enzyme and substrate (Fersht, 1985); perhaps we should wonder why the cellular levels of the diphosphoinositol polyphosphates are actually as high as they are. All of the DIPPs are relatively small proteins of just under 20 kDa. The active site of each is based on the so-called Nudix motif, which is typically, although not exclusively, Gx5Ex5[UA]xREx2EExGU (U represents an aliphatic, hydrophobic residue) (McLennan, 2006). The DIPPs represent an unusual context in which to find this motif; it is more usually reserved for proteins whose functions are limited to the hydrolysis of nucleoside di- and triphosphates, nucleotide sugars, and dinucleoside polyphosphates (McLennan, 2007). A detailed mutagenic study has revealed that the specificity of human DIPP1 toward diphosphoinositol polyphosphates is entrusted to several amino acid residues that lie outside the Nudix motif (Yang et al., 1999). Interestingly, a Nudix hydrolase from African Swine Fever Virus (g5R) has also been shown to have considerable DIPP activity, and so it was suggested that a reduction in cellular levels of diphosphoinositol polyphosphates might be important for viral morphogenesis (Cartwright et al., 2002). The Nudix motif is also utilized by mRNA decapping enzymes (Dunkley and Parker, 1999; Parrish et al., 2007). As a consequence, it was recently speculated that decapping might in some cases be regulated by active-site competition between diphosphoinositol polyphosphates and capped mRNA substrates (McLennan, 2007). However, we have found that two mRNA decapping enzymes with Nudix motifs, X29 and DCP2, do not dephosphorylate diphosphoinositol polyphosphates (Cho, Kiledjian, Shears, unpublished data). The a- and b-isoforms of hDIPP2 are distinguished solely by the latter having an additional glutamine residue, but both are encoded by a single gene with an AGCAG pentamer that offers two adjacent, alternate intronic 30 -boundaries. Thus, “intron boundary skidding” (Caffrey and Shears, 2001) was introduced as a term to describe this mechanism for yielding both DIPP2a and hDIPP2b mRNAs from a single gene. The b-isoform has about one third of the activity of the a-isoform (Caffrey and Shears, 2001). As for the a- and b-isoforms of hDIPP3, these also differ by only one amino-acid residue: Pro-89 in hDIPP3a is replaced by an Arg in hDIPP3b. This substitution makes the b-form about 2.5-fold more active (Hidaka et al., 2002). These two proteins are separately encoded by duplicated genes on chromosome X (Hidaka et al., 2002; Leslie et al., 2002). Transcription of both genes is inactivated on one of the X chromosomes of human females to maintain appropriate gene dosage (Hidaka et al., 2002). This geneduplication event occurred prior to the divergence of primates and sciurognath rodents, approximately 115 million years ago (Hua et al., 2003), greatly exceeding the 4-million-year half-life for inactivation of redundant paralogues (Lynch and Conery, 2000). Since the DIPP3 duplication has been maintained for much longer, it must be functionally significant, perhaps facilitating tissue-dependent expression patterns (Hua et al., 2003). Not only are there multiple DIPP proteins, but, especially for the DIPP2s (Caffrey et al., 2000), there is evidence of considerable alternate splicing within the UTRs. The significance of there being multiple mRNAs is typically interpreted in terms of their differing in either stability or translatability, to modulate gene expression in a tissue- or developmental stage-specific manner (Edwalds-Gilbert et al., 1997). There are some clues within their sequences that the DIPP mRNAs may turn over rapidly: Alu repeat sequences in the DIPP2 mRNA (Caffrey et al., 2000) and Type 1 AU-rich elements in the DIPP3 mRNAs (Hidaka et al., 2002). This possibility has not yet been tested experimentally, but it remains the most promising route for unearthing factors that might regulate cellular DIPP activity; there is no evidence as yet that the inherent activities of the enzymes are regulated. To date, the experimental manipulation of the expression of the DIPPs has yielded only a limited amount of biological information. Schizosaccharomyces pombe showed no obvious phenotype following disruption of the Aps1 gene (which encodes a DIPP homologue) (Ingram et al., 2003). However, it is of interest that the diphosphate species that increased in levels the most in the aps1D cells was a generally barely detectable compound tentatively identified as [PP]2-InsP3 (the twice diphosphorylated derivative of Ins(1,3,4,5,6)P5, see > Figure 12-1). Considering that levels of Ins(1,3,4,5,6)P5 are so low in yeast cells,

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it was suggested that the metabolic flux between Ins(1,3,4,5,6)P5 and [PP]2-InsP3 must normally be exceptionally high (Ingram et al., 2003). This is a prescient conclusion; 4 years later, “hidden” but metabolically active pools of Ins(1,3,4,5,6)P5 (and InsP6) were also proposed to exist in mammalian cells (Otto et al., 2007). The functions of the diphosphoinositol polyphosphates were also not illuminated by overexpression of Aps1. Although these cells became swollen, rounded, and multiseptate, the relationship of these effects to the diphosphoinositol polyphosphates is unclear, since, surprisingly, cellular levels of these polyphosphates were unaffected by overexpressing Aps1 (Ingram et al., 2003).

4

Proposed Mechanisms of Action of Diphosphoinositol Polyphosphates

4.1 A Novel Mode of Posttranslational Modification It has long been recognized that the removal of one of the highly polar phosphate groups from a diphosphoinositol polyphosphate offers the molecule significant relief from the severe electrostatic and steric congestion around the inositol ring; the standard free energy change during their hydrolysis must be quite substantial (Stephens et al., 1993; Laussmann et al., 1996; Hand and Honek, 2007). Snyder and colleagues have, for some time (e.g., see Luo et al., 2002), been investigating the possibility that this free energy change might enable diphosphoinositol polyphosphates to phosphorylate proteins. Their data were first published in 2004 (Saiardi et al., 2004). The in vitro results are very striking: PP-InsP5 can donate the b-phosphate in the diphosphate group to several nucleolar proteins, including Nsrl (yeast nucleolin), Nopp140, and TCOF1 (Saiardi et al., 2004). The common phosphorylation site in each of these target proteins was a serine that is surrounded by acidic residues. Although this study indicated that PP-InsP5 functioned completely independent of protein kinases (Saiardi et al., 2004), it emerged soon afterward that PP-InsP5 phosphorylates its targets only after they are first “primed” by an initial casein kinase 2 (CK2)-dependent phosphorylation event (York and Hunter, 2004; Bennett et al., 2006; Bhandari et al., 2007). Furthermore, the evidence now points to the serine that is phosphorylated by CK2 subsequently being converted to a diphosphate by PP-InsP5 (Bhandari et al., 2007). These data provide a provocative and novel idea concerning the molecular action of diphosphoinositol polyphosphates while also unveiling evidence of a novel mechanism of covalent modification: diphosphorylation of serine. But, does this occur in vivo? And, if so, is the extent of posttranslational modification driven by fluctuations in the cellular levels of the diphosphoinositol polyphosphates? Such a regulatory paradigm is constrained by there being only a few known biologically relevant stimuli that can elicit significant changes in the levels of diphosphoinositol polyphosphates (> Section 3.1). To examine if there is any relationship between cellular levels of diphosphoinositol polyphosphates and Nsr1 phosphorylation, Snyder and colleagues (2004) used yeast cells in which the InsP6 kinase (Kcs1) that makes 5-PP-InsP5 was genetically eliminated. They found that the degree of phosphorylation of endogenous Nsr1 was substantially reduced in these cells. This is a promising observation, but it should be noted that the deletion of Kcs1 impairs a number of biological processes in yeasts: vacuolar biogenesis (Saiardi et al., 2000; Saiardi et al., 2002), endocytosis (Saiardi et al., 2002), stress responses (Dubois et al., 2002), and DNA hyperrecombination (Luo et al., 2002). In such circumstances, it is not unlikely that an effect upon Nsr1 phosphorylation could arise independent of PPInsP5 synthesis. Arguably of more interest is the observation that the deletion of the Nsr1 gene caused a doubling of intracellular levels of PP-InsP5 and [PP]2-InsP4 (Saiardi et al., 2004). The inference here is that Nsr1 phosphorylation is an ongoing process that must normally consume considerable quantities of the diphosphoinositol polyphosphates. If this assessment is correct, then the rate of turnover of diphosphoinositol polyphosphates in vivo must be even more astonishing than we currently blame on the high activity of DIPPs (> Section 3.2). However, once again, we should be cautious: the changes in levels of PP-InsP5 and [PP]2-InsP4 in the nsr1D strain might be a secondary consequence of the accompanying growth-impaired phenotype. Another potential obstacle that might prevent protein phosphorylation by PP-InsP5 from being a physiologically relevant process is that this reaction is inhibited by InsP6 (Saiardi et al., 2004) that is present

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in cells at 25-fold higher levels than is PP-InsP5. One can always argue that this impediment can be circumvented by compartmentalization of InsP6 away from PP-InsP5 and its target. In fact, there are reasons to believe that at least a portion of the cell’s InsP6 pool might be immobilized by its binding to membranes (Poyner et al., 1993) or to proteins (Shears, 2001). Additionally, data in a very recent report indicate that cells may contain pools of InsP6 that show different metabolic characteristics (Otto et al., 2007). Physical compartmentalization could explain the latter phenomenon. Indeed, data showing a punctate distribution of the Ins(1,3,4,5,6)P5 2-kinase within certain cellular structures such as nucleoli and stress-granules strongly indicate that intracellular InsP6 synthesis is compartmentalized (Brehm et al., 2007). Thus, one can turn around the argument that InsP6 might be an “obstacle” (see earlier), and instead suggest a potential regulatory mechanism whereby dynamic, localized changes in the extent to which InsP6 inhibits protein phosphorylation by diphosphoinositol polyphosphates. One might also presume that the reverse reaction—dephosphorylation of the protein—might also be a regulated event. So far, a specific phosphatase that might achieve this has not been identified. In fact, the putative diphosphorylated protein seems quite resistant to enzymatic hydrolysis (Bhandari et al., 2007). Finally, there is either a blessing and a curse—depending upon your point of view—in the demonstration that not just 5-PP-InsP5, but also the other isomer of PP-InsP5 made by yeast Vip1 (> Section 3.1), and [PP]2-InsP4, can all phosphorylate the same proteins (Bhandari et al., 2007). The good news one might take from this observation is that the biological significance of diphosphoinositol polyphosphates is greatly expanded. But, the bad news is this: it does not easily explain the selectivity of some of the biological actions of the different diphosphorylated molecules that are clearly demonstrated in the literature (> Section 5).

4.2 Competitive Ligand Binding Immediately following the initial characterization of the diphosphoinositol polyphosphates (Menniti et al., 1993; Stephens et al., 1993), my laboratory searched for proteins that bound these molecules in the hope that we would discover a biologically informative “receptor.” This screen uncovered several proteins that all have in common a role in regulating vesicular traffic: Coatomer, AP-2, and AP180 (previously sometimes called “AP-3”) each bind PP-InsP5 with high affinity (Fleischer et al., 1994; Shears et al., 1995; Ye et al., 1995). Subsequently, we defined the ligand-binding site of AP180 in more detail, and we found that inositol lipids bound to the same site as PP-InsP5 (Hao et al., 1997). However, PP-InsP5 binds with a 35-fold higher affinity than does PtdIns(3,4,5)P3 (Ye et al., 1995; Hao et al., 1997). This has raised the possibility that there may be functional significance to this ligand competition, although this has yet to be put into any biologically meaningful context. This ligand-binding site was initially labeled as an epsin N-terminal homology (ENTH) domain, but it later emerged as a distinct—and eponymous—ANTH (AP180 N-terminal homology) domain (see Ford et al., 2001; Legendre-Guillemin et al., 2004). The ANTH ligand-binding site is unusual in being surface exposed, rather than in a pocket or groove (Ford et al., 2001). The phosphates are perched on the tips of the side chains of three lysines and a histidine; this arrangement has been likened to a ball balanced on the fingertips (Ford et al., 2001). Clearly, the inositol headgroup of either PtdIns(4,5)P2 or PtdIns(3,4,5)P3 has a very different spatial arrangement of phosphate groups compared with the diphosphoinositol polyphosphates. Nevertheless, there are two factors that enable these two classes of ligands to successfully compete for the same protein domain. First, the ANTH-binding site is surface mounted, limiting any topographical constraints. Second, delocalized electrostatic interactions of a protein with a multiphosphorylated molecule (such as PP-InsP5) can substitute for specific ligand interactions driven by a geometrically precise arrangement of fewer phosphate groups (Lemmon et al., 2002). However, these same ligand-binding characteristics leave little room for discrimination between different isomers of PP-InsP5. Thus, just as is the case with protein phosphorylation by diphosphoinositol polyphosphates (> Section 4.1), this competitive ligand-binding scenario does not explain the structural specificity of some biological actions of the different diphosphorylated molecules (> Section 5).

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Some PH domains also bind ligands through delocalized electrostatics (Lemmon et al., 2002). Snyder and colleagues (Luo et al., 2003) have reported that PP-InsP5 competes with PtdIns(3,4,5)P3 for binding to the PH domain of Dictyostelium CRAC (cytosolic regulator of adenylyl cyclase) (Luo et al., 2003). The relative affinities of these two competing ligands were not directly quantified, although PP-InsP5 was reported to be approximately equipotent with Ins(1,3,4,5)P4, the headgroup of PtdIns(3,4,5)P3 (Luo et al., 2003). Evidence was provided that this competition between PP-InsP5 and PtdIns(3,4,5)P3 regulated the intracellular distribution of the protein, which is a key factor for determining the directionality of chemotaxis of Dictyostelium (Luo et al., 2003). However, compared with Dictyostelium, the levels of diphosphoinositol polyphosphates in mammalian cells are up to 100-fold lower, making it more difficult for PH domains in higher organisms to utilize PP-InsP5 as a physiologically relevant competitor with PtdIns (3,4,5)P3. Moreover, Downes and colleagues (Komander et al., 2004; Downes et al., 2005) have been unable to reproduce the observation of the Snyder laboratory (Luo et al., 2003) that PP-InsP5 has such a high affinity for PH domains. Perhaps the origin of these quantitative experimental differences between the two laboratories lies in the way that each group prepared and quantified their PP-InsP5.

5

Biological Effects

A number of biological effects have been blamed upon the diphosphoinositol polyphosphates. Some of these ideas arose from genetic disruption of the turnover of diphosphoinositol polyphosphates, by deletion of Ksc1 for example (see later). The possibility of unexpected side effects should always be considered as a potential caveat in such experiments. For example, PtdIns(4,5)P2 levels were, by an unknown mechanism, reduced by nearly 40% in the kcs1D strain (Saiardi et al., 2002). Considering how multifunctional is PtdIns(4,5)P2, a change in its levels could cause any number of phenotypic changes. Moreover, DNA microarrays and Northern analyses have identified 30 genes in S. cerevisiae, the expression of which was influenced by Kcs1 (El Alami et al., 2003). Thus, the phenotype of the kcs1D strain might not be a direct consequence of the deletion of Kcs1, but instead it might result from these secondary genetic effects.

5.1 DNA Maintenance and Repair Even before Kcs1 from S. cerevisiae was shown to encode InsP6 kinase activity, it was known to promote recombination in yeast strains with defective PKC activity (Huang and Symington, 1995). Subsequently, impaired homologous recombination in the kcs1D strain was shown to underlie a reduced ability to repair damaged DNA (Luo et al., 2002). To date, this area of research has not been pursued further. In a separate development, two groups working independently—Snyder’s laboratory at Johns Hopkins (Saiardi et al., 2005) and York’s team at Duke University (York et al., 2005)—reported that some of the diphosphoinositol polyphosphates regulate another aspect of DNA integrity, namely, telomere length. Telomeres are the nucleoprotein complexes that occur at the ends of eukaryotic linear chromosomes. They consist of long, repetitive DNA sequences that attract a number of sequence- and structure-specific binding proteins (for reviews, see d’Adda et al., 2004; Smogorzewska and de Lange, 2004). These chromosomal caps prevent nucleolytic degradation and provide a mechanism for cells to distinguish natural termini from broken ends; the latter signal DNA damage. Without telomere protection, the activation of DNA damage response pathways by chromosomal ends would provoke cell cycle arrest, senescence, or apoptosis. Another feature of telomeres is that their 30 -single-strand nucleotide overhang cannot be replicated, so most human somatic cells lose terminal DNA with each division; thus, telomeres also protect the more internally located coding region of the genome, but this protection requires ongoing telomere attrition to be countered by elongation mechanisms. The Snyder and York teams found that telomere length was inversely proportional to the degree of Kcs1 expression (Saiardi et al., 2005; York et al., 2005). For example, elimination of the diphosphoinositol polyphosphates, by deletion of the Kcs1 gene, was accompanied by telomere lengthening (Saiardi et al., 2005; York et al., 2005). However, Kcs1 phosphorylates both Ins(1,3,4,5,6)P5 and InsP6

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(Saiardi et al., 2000; > Figure 12-1). To determine if the diphosphates of either Ins(1,3,4,5,6)P5 or InsP6 (or both) regulated telomere length, the Ins(1,3,4,5,6)P5 2-kinase gene was deleted (see > Figure 12-1). This maneuver resulted in an increased phosphorylation of Ins(1,3,4,5,6)P5 to PP-InsP4, which in turn was associated with decreased telomere length (Saiardi et al., 2005; York et al., 2005). York et al. (2005) added an additional experiment that is critical in view of the pleiotropic nature of Kcs1 (Dubois et al., 2002): while transformation of kcs1D yeast with wild-type Kcs1 rescued telomere length, transformation with a kinasedead mutant of Kcs1 (Asp791Ala; Lys793Ala) did not. These results specifically attribute telomere maintenance to the kinase activity of Kcs1 and not to some additional domain in this protein. Both groups suggest that a fluctuating PP-InsP4 concentration acts as a rheostat that can either extend or shorten telomeres. Among the important players in telomere maintenance in yeast are Tel1 and Mec1 (Mallory and Petes, 2000), members of the family of phosphoinositide 3-kinase related kinases (PIKKs). Despite homology in their catalytic domain to phosphatidylinositol 3-kinases, the PIKKs phosphorylate proteins rather than lipids. Tel1, at least, appears to be downstream of Kcs1 (Saiardi et al., 2005; York et al., 2005). Perhaps PP-IP4 is a negative regulator of Tel1 and/or Mec1. Indeed, in their study, Snyder’s group (Saiardi et al., 2005) asserted that they “. . .have shown that inositol pyrophosphates [sic] physiologically inhibit signaling by Tel1,” but this was a rather premature claim that was not actually directly validated. The mechanism of action of PP-InsP4 has not yet been determined.

5.2 Yeast Vacuole Biogenesis and Vesicle Trafficking AP180 is a brain-specific protein that binds clathrin and helps assemble it into cages during the initial stages of synaptic vesicle recycling (Legendre-Guillemin et al., 2004). In an in vitro assay, we found that PP-InsP5 inhibited AP180-mediated clathrin cage assembly (Ye et al., 1995). This suggests that PP-InsP5 might inhibit endocytosis. Consistent with this idea, it has been reported that endocytosis was accelerated in the kcs1D strain of S. cerevisiae (Saiardi et al., 2002). This apparently accounts for the formation of small, fragmented vacuoles in the kcs1D cells, rather than the single large vacuole that is typical of wild-type yeast (Saiardi et al., 2000, 2002; Dubois et al., 2002). [PP]2-InsP4 binds to AP180 at a so-called ANTH domain, which also binds inositol lipids (> Section 4.2). Most of the current literature focuses on the inositol lipids as being the biologically significant ligands, and the effects of the diphosphoinositol polyphosphates upon AP-180 have largely been ignored (Legendre-Guillemin et al., 2004). Moreover, the consensus is that the role of lipid binding, most notably for PtdIns(4,5)P2, is to promote endocytosis (Legendre-Guillemin et al., 2004). This has rather muddied the waters with regard to understanding the significance of the in vitro clathrin assembly assays, which indicated that both the lipids (Hao et al., 1997) and the diphosphoinositol polyphosphates (Ye et al., 1995) would inhibit endocytosis. Nevertheless, it remains possible that there is physiological significance to this competition between these two sets of ligands, both with regard to AP180 function and that of other proteins with ANTH-domains.

5.3 Apoptosis The idea that diphosphoinositol pyrophosphates might play a role in apoptosis originates with the demonstration that an antisense-mediated “knockdown” of IP6K-type 2 expression prevented the apoptotic actions of interferon-b in ovarian carcinoma cells (Morrison et al., 2001, 2002). Conversely, overexpression of IP6K-type 2 augmented apoptosis (Morrison et al., 2001, 2002). In a later study (Nagata et al., 2005), overexpression of either of the three mammalian IP6Ks was shown to augment the cytotoxic actions of several cell stressors, including hypoxia, hydrogen peroxide, and also cisplatin, a widely used antineoplastic agent the cytotoxicity of which is apparently due to it forming DNA adducts (Choi and Kim, 2006). Catalytically dead InsP6 kinase did not promote apoptosis (Nagata et al., 2005). The conclusion that a rise in PP-InsP5 levels might be pro-apoptotic was reinforced by the further observation that cisplatin treatment by itself increased the cellular concentration of PP-InsP5

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(Nagata et al., 2005). In further experiments, the endogenous expression of each of the three kinases was individually targeted by RNAi. A reduction in IP6K-type 2 expression protected the cell against the actions of apoptotic stimuli (Nagata et al., 2005). Knockdown of the type 1 and type 3 isoforms did not mimic this effect (Nagata et al., 2005). The reason why the type 2 kinase has this unique effect may be associated with its stress-dependent relocation from the nucleus to Bax-positive (i.e., damaged) mitochondria; the type 1 and 3 kinases do not show this effect (Nagata et al., 2005). This particular observation suggests that apoptosis can be accelerated by an elevated synthesis of PP-InsP5 near the mitochondria. The authors further suggested that under physiological conditions the type 2 kinase might serve this function, but following overexpression of any of the InsP6 kinases, a global elevation in PP-InsP5 levels seems adequate. However, it should be noted that a subsequent study failed to describe stress-dependent relocation of the type 2 InsP6 kinase to mitochondria during apoptosis (Morrison et al., 2005). The latter study instead indicated that it is the migration of the type 2 kinase into the nucleus that promotes apoptosis (Morrison et al., 2005). Clearly, further work on this topic is required to unravel the mechanistic basis for this apparent interrelationship between PP-InsP5 synthesis and apoptosis.

5.4 Transcriptional Regulation In S. cerevisiae, the expression of Pho11 and Pho5 acid phosphatases, and some phosphate transporters, is controlled by Pho4, a transcriptional regulator that is phosphorylated by the Pho80-Pho85 cyclin/cyclindependent kinase complex (Kaffman et al., 1994). When cells are grown on phosphate-rice media, the cyclin kinase complex is active, Pho4 is phosphorylated, and exported out of the nucleus; when phosphate supply is limited, the cyclin kinase complex is inactive, and dephosphorylated Pho4 becomes concentrated in the nucleus and activates transcription of the phosphate-responsive genes (Kaffman et al., 1994). But, how does the cell sense change in extracellular phosphate availability? An early indication that diphosphoinositol polyphosphates might be involved in this process arose from the finding that the expression of the Pho11 acid phosphatase is inversely correlated with changes in Kcs1 (InsP6 kinase) expression (El Alami et al., 2003). Later work saw a similar, negative correlation with regard to Pho5 expression; the expression of this acid phosphatase was constitutively upregulated in kcs1D yeast (Auesukaree et al., 2005). These studies suggested that the expression of acid phosphatases in yeast might be regulated by one or more diphosphoinositol polyphosphates. However, despite these early experiments indicative of a role for Kcs1, a recent report indicates that Ppip5k/Vip1 is more important. O’Shea and colleagues (Lee et al., 2007) discovered that reconstitution in vitro of the inhibition of the Pho80–Pho85 kinase complex by Pho81 required an essential cofactor: the PP-InsP5 isomer made by Ppip5k/Vip1. Moreover, it was found that Pho4 was constitutively cytoplasmic in a vip1D strain of S. cerevisiae, even in phosphate-starved yeast, suggesting that the Pho80–Pho85 cyclin/cyclin-dependent kinase complex could not then be inactivated (Lee et al., 2007). In kcs1D cells, Pho4 could be dephosphorylated and thereby enter the nucleus as normal. Overall, it seems to be that cyclin kinase activity is regulated by the novel PP-InsP5 made by Vip1/Ppip5k (i.e., the proposed 1/3-diphosphate; see > Section 3.1), and not the 5-PP-InsP5 made by Kcs1 (Lee et al., 2007). These data speak to a novel mode of action of a diphosphoinositol polyphosphate, since current mechanistic proposals do not include an explanation for such specificity of action (> Section 4). Indeed, recent studies (Lee et al., 2008) indicate that, through non-covalent binding to Pho81, 1/3-PP-InsP5 modifies the interactions between Pho81 and Pho-80/ Pho85, thereby regulating the access of substrates to the cyclin kinase active site. This latest development offers a classical protein-ligand interaction as the mechanism of action that is more typical for second messengers. These observations were placed in a regulatory context by O’Shea’s HPLC data (Lee et al., 2007), which showed that limitation of extracellular phosphate supply (i.e., its reduction to 10 mM Pi) prompted a tenfold increase in the levels of two [3H]inositol-labeled compounds more polar than InsP6. These two molecules could be PP-InsP5 and [PP]2-InsP4, or possibly the two different PP-InsP5 isomers now known to exist in cells (> Figure 12-1). One of these two HPLC “peaks” was proposed to be the PP-InsP5 isomer that inhibits the cyclin kinase complex.

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This is a fascinating development, although there are aspects of this phenomenon that could benefit from some further clarification: An IC50 of 55 mM was obtained for the 1/3-PP-InsP5-facilitated inhibition of cyclin kinase activity in vitro (Lee et al., 2007). O’Shea and colleagues (Lee et al., 2007) conclude that this is physiologically relevant because they estimate that total PP-InsP5 levels rise to 20–30 mM in yeast limited to 10 mM Pi. However, there are reasons to doubt that calculation. No direct mass measurements of 1/3-PPInsP5 concentration were made. Instead, they estimate total intracellular PP-InsP5 concentration from the ratio, PP-[3H]InsP5/[3H]InsP6, by assuming that the InsP6 level is 100 mM. This could turn out to be an overoptimistic estimate; there is one report of a mammalian cell line containing this much InsP6 (Bunce et al., 1993), but estimates for other cells are in the 15–60 mM range (Szwergold et al., 1987; Pittet et al., 1989; Irvine and Schell, 2001; Barker et al., 2004), and there are even reasons to think 60 mM might be the upper limit of InsP6 solubility (Irvine and Schell, 2001; Torres et al., 2005). Additionally, the calculation also assumes that InsP6 levels do not themselves increase during phosphate limitation, which appears not to be correct (Lee et al., 2007). Moreover, the estimate of total PP-[3H]InsP5 is derived from the two more-or-less equally sized [3H]inositol-labeled peaks that elute after InsP6 during HPLC analysis (Lee et al., 2007). Only one half of this total can actually represent 1/3-PP-InsP5, the only PP-InsP5 isomer that was inhibitory in the reconstituted cyclin kinase assays (see earlier). Overall, therefore, it can be argued that there is a considerable disparity between the efficacy of 1/3-PP-InsP5 action in vitro and its cellular levels in vivo. This is an issue that deserves further study. It is also arguable that a rise in PP-InsP5 levels in response to a loss of extracellular phosphate is a somewhat counter-intuitive signaling response. One might have anticipated that cells would conserve ATP in such a situation, rather than increase its rate of utilization through the increased synthesis of diphosphoinositol polyphosphates. In fact, a different laboratory has stated that they are unable to repeat the observation that PP-InsP5 synthesis is increased in yeast under conditions of limited phosphate supply (Onnebo and Saiardi, 2007). However, we should wait for these purportedly contradictory data to be published before we draw any firm conclusions.

5.5 Exocytosis A recent study has explored the effect of PP-InsP5 upon the size of the rapidly releasable pool of insulin vesicles in pancreatic b-cells (Illies et al., 2007). This subset of insulin granules is in close proximity with the plasma membrane, ready to undergo exocytosis as soon as the cell receives the appropriate stimulus (e.g., glucose). Overexpression of the InsP6 kinase more than doubled the size of this readily releasable pool, according to single-cell capacitance measurements (cell capacitance increases as the cell surface area is enlarged, following the fusion of exocytic vesicles). A catalytically dead kinase construct did not imitate this effect, illustrating that the increased exocytosis was dependent upon PP-InsP5 synthesis (Illies et al., 2007). Similar effects were observed using a human growth hormone secretion reporter assay for exocytosis (Illies et al., 2007). Moreover, the initial phase of insulin secretion could also be enhanced by the direct application of PP-InsP5, within a physiologically relevant range of concentrations (1–10 mM), whereas InsP6 had no effect (Illies et al., 2007). Conversely, insulin secretion was reduced after endogenous PP-InsP5 levels were attenuated by the knockdown of InsP6 kinase by RNAi (Illies et al., 2007). Interestingly (from the point of view of specificity of action) the rate of exocytosis of the reserve pool of insulin granules was not affected by PP-InsP5 (Illies et al., 2007). Although these observations are impressive in themselves, there is also now excitement for the possibility that PP-InsP5 might also stimulate other exocytic processes, such as neurotransmitter release, for example. There is no information yet concerning the possible mechanism by which PP-InsP5 stimulates exocytosis. The phenomenon is not specific for the isomer with 5-diphosphate; several other isomers of PP-InsP5 were as efficacious as 5-PP-InsP5 (Illies et al., 2007). Molecular mechanisms that do not discriminate between different PP-InsP5 isomers include protein phosphorylation and competitive binding with inositol lipids (> Section 4). Either could play a role in this process. The race is now on to identify the PP-InsP5 target.

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PTEN and PI3 Kinase Signaling in the Nervous System

C. P. Downes . B. J. Eickholt . M. L. J. Ashford . N. R. Leslie

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246 1.1 PI3K Generates Two Lipid Signals with Prominent Roles in Neuronal Development and Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246 1.2 Class I PI3K Signaling is Antagonized by PTEN . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 249 1.3 The PTEN Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 249 2 2.1 2.2 2.3 2.4

PI3K/PTEN Signaling Controlling Neuronal Morphogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 250 Breaking Symmetry – PI3K/PTEN Signaling During Neuronal Polarization . . . . . . . . . . . . . . . . . . . . . 250 Spatially Restricted PIP3 Accumulations Regulate Cell Polarity in Nonneuronal Cells . . . . . . . . . . . . 250 PI3K/PTEN Signaling During Extension and Navigation of Neurites . . . . . . . . . . . . . . . . . . . . . . . . . . . . 252 Neurite Branching and Dendritic Morphogenesis are Controlled by PI3K/PTEN Signaling . . . . . 252

3 3.1 3.2 3.3 3.4

PI3K/PTEN Signaling in Normal Brain Function and Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 253 Learning and Memory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 253 Seizure and Epilepsy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 253 PTEN/PI3K Signaling and Autism Spectrum Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 254 PTEN is Required for Addictive Responses to Drugs of Abuse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255

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PI3K/PTEN Signaling in the Central Control of Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 255

5 5.1 5.2 5.3

PI3K/PTEN Signaling and Neurodegenerative Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256 Ischemic Brain Injury/Stroke . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256 Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 257 Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 258

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Future Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 259

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Abstract: Class I phosphoinositide 3-kinases (PI 3-kinases) synthesise the lipid second messenger, phosphatidylinositol 3,4,5-trisphosphate (PIP3) in response to growth factors, neurotransmitters, hormones, cell-cell and cell-matrix contacts. This signal in turn is either removed by the tumour suppressor, phosphatase and tensin homologue deleted on chromosome ten (PTEN), or further metabolised by 5-phosphatases to generate another signal lipid, phosphatidylinositol 3,4-bisphosphate (PI(3,4)P2). PIP3 and PI(3,4)P2 initiate complex signalling cascades through their interactions with highly specific lipid binding domains, most commonly pleckstrin homology (PH) domains, present within a broad range of target proteins. This so-called PI 3-kinase signaling pathway is prominent in both the developing and mature nervous systems of mammals as well as less complex organisms. This review focuses on the mutually antagonistic roles of class I PI 3-kinases and PTEN both of which are required for the development of cell polarity, an essential aspect of neuronal development and morphogenesis as well as neurite extension and navigation. In addition we discuss the importance of this system in diseases of the nervous system. Included in the latter is a survey of the mounting evidence that PI 3-kinase/PTEN signaling may be important in autism spectrum disorders, addictive responses to drugs of abuse, diseases such as diabetes and obesity involving, in part, the central control of metabolism, and several of the most common neurodegenerative diseases. List of Abbreviations: APP, amyloid precursor protein; ASD, autistic spectrum disorders; BACE1, beta site APP-cleaving enzyme; BAD, Bcl-2 associated death protein; PI3Ks, phosphoinositide 3-kinases; PI(4,5)P2, phosphatidylinositol 4,5-bisphosphate; PIP3, phosphatidylinositol 3,4,5-trisphosphate; PKB, protein kinase B; POMC, proopiomelanocortin; PTEN, phosphatase and tensin homologue deleted on chromosome ten; ROS, reactive oxygen species; TSC, tuberous sclerosis complex; VTA, ventral tegmental area

1

Introduction

The inositol glycerophospholipids are amongst the most versatile and ubiquitous classes of signaling molecules in eukaryotic cells. Their functions range from the generation of multiple second messengers in response to hormones, neurotransmitters, and growth factors and the control of cell shape via regulation of the actin cytoskeleton, to participation in membrane trafficking dynamics by functioning as specific postmarks of particular subcellular compartments. Each of these functions has been adapted to the specific needs of the nervous system at all stages of its development and in the adult. This review focuses on one important aspect of phosphoinositide signaling, namely the signaling cascades initiated by activation of Class I Phosphoinositide 3-kinsases (PI3Ks) and their reversal by the tumor suppressor, phosphatase and tensin homologue deleted on chromosome ten (PTEN) with particular emphasis on their roles in the establishment and maintenance of a polarized cell state and in diseases of the nervous system. PTEN is primarily seen as an antagonist of PI3K signaling, but it is important to reconsider this understandable paradigm. Cell polarity and related phenomena, such as directed cell movement, require the generation of relatively stable signal gradients which can only be achieved by the coordinated spatial and temporal regulation of both synthetic and catabolic processes. This review, therefore, emphasizes recent work describing both the subcellular targeting and negative regulation of PTEN, which appear necessary for some PI3K-dependent biological responses.

1.1 PI3K Generates Two Lipid Signals with Prominent Roles in Neuronal Development and Function Class I PI3Ks synthesize the lipid second messenger, phosphatidylinositol 3,4,5-trisphosphate (PIP3) from its relatively more abundant precursor, phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) (Vanhaesebroeck et al., 2001). These enzymes are heterodimers comprising of one of four catalytic subunits (a, b, g, d) tightly bound to a regulatory subunit. The a, b, and d isoforms combine with one of five regulatory proteins (p85a, p55a, p50a, p85b, or p55g) which convey sensitivity to tyrosine kinase-dependent signaling pathways, since each contains a pair of phosphotyrosine-binding SH2 domains. By contrast, the g isoform of

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PI3K is complexed with 101 kDa or 84 kDa regulatory domains which convey high sensitivity to activation by the bg-subunits of heterotrimeric G-proteins. In addition, all the catalytic subunits have Ras-binding motifs which may account for the ability of activated forms of Ras to trigger PI 3-kinase-dependent signaling pathways. Hence PI3K signaling can be initiated by a broad range of input signals, including growth factors, insulin, cytokines, neurotrophic factors, extracellular matrix components and in response to activation of several G-protein coupled receptors. All of the Class I PI3K catalytic and regulatory subunits are expressed in both the developing and mature mammalian nervous system. EST database information from the adult human brain suggests an expression level of approximately 40 transcripts per million for p110b, with p110a and p110d being approximately fivefold lower, and p110g another fivefold lower still. However, it is interesting to note that the expression of p110b and p110d PI3K in the mouse brain is enriched relative to other tissues and that p110d is expressed almost exclusively in the liver, hematopoietic system, and the developing nervous system at both embryonic and adult stages (Eickholt et al. 2007). PIP3 can be metabolized by members of the inositol polyphosphate 5-phosphatase family (eg SH2 domain containing inositol 5-phosphatases 1 and 2, SHIP1 and SHIP2) to generate a second output signal of Class I PI3Ks, namely phosphatidylinsoitol 3,4-bisphosphate (PI(3,4)P2). PIP3 and PI(3,4)P2 bind to overlapping sets of effector proteins each of which possesses a selective lipid binding domain, most commonly a pleckstrin homology (PH) domain (Lemmon, 2003) (See > Figure 13-1). This contributes to the assembly of signaling complexes at the plasma membrane and in some cases also leads to activation of the effector itself. One of the best studied examples illustrated is the protein kinase Akt (also known as

. Figure 13-1 Model for the inhibition of PI 3-Kinase signaling by PTEN and 5-phosphatases. PIP3 is synthesized at the plasma membrane from the abundant cellular phosphoinositide PI(4,5)P2 by the regulated action of class I PI3K enzymes. The localization and abundance of the 3-phosphorylated lipid products is also affected by two classes of phosphatases: the 3-phosphatase PTEN converts PIP3 back to PI(4,5)P2 and the phosphoinositide 5-phosphatases, exemplified by the SHIP proteins and SKIP, convert PIP3 into PI(3,4)P2. PIP3 and PI(3,4)P2 mediate their effects on downstream signaling and cellular behavior through protein targets that are able to recognize these lipids and bind them selectively. Each lipid has unique targets, such as GRP1 and P-Rex-1 for PIP3 and the TAPP proteins and lamellipodin for PI(3,4)P2, but they also share several targets, such as Akt and DAPP1, that appear to bind to both lipids and mediate signaling upon the generation of either lipid

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Protein Kinase B, or PKB) which possesses a PH domain that binds either PIP3 or PI(3,4)P2 (Lawlor and Alessi, 2001; Fayard et al., 2005). Activation of PI3K causes translocation of Akt to the plasma membrane and a conformational change that makes two residues (Thr 308 and Ser 473) available for phosphorylation by other protein kinases. One of these, phosphoinositide dependent kinase 1 (PDK1) has a PH domain with a similar lipid specificity and thus is colocalized with Akt accounting for phosphorylation of the latter at Thr 308. Recent studies suggest that a complex containing mammalian target of rapamycin (mTOR) and Rictor probably accounts for phosphorylation of PKB at Ser 473 (Sarbassov et al., 2005). In addition, several guanine nucleotide exchange factors for Rho family GTPases which control cell shape and motility, have PH domains that bind only PIP3 with high affinity (Welch et al., 2003) and others, such as the TAPP proteins and lamellipodin appear to bind only PI(3,4)P2 (Dowler et al., 2000; Krause et al., 2004). Although these and other molecules have been identified as bona fide direct targets of PIP3 or PI(3,4)P2, other molecules, including several ion channels, have been proposed to be targets for PIP3 regulation (Le Blanc et al., 2004; Brady et al., 2006); however, data are currently lacking regarding the sufficiently selective binding of these proteins to PIP3 to support them as direct targets, and some evidence indicates indirect regulation of some ion channels through PIP3-dependent plasma membrane recruitment or phosphorylation downstream of Akt (Viard et al., 2004; Sun et al., 2006). These and other signaling events downstream of PI3K which have particular relevance to the nervous system are illustrated in > Figure 13-2. In the developing nervous system they provide key contributions to

. Figure 13-2 Downstream targets of PIP3. Several downstream targets of PIP3 have been shown to bind directly and selectively to this lipid. Direct interactions are shown as solid arrows (Vanhaesebroeck and Alessi, 2000; Vanhaesebroeck et al., 2001; Lemmon, 2003; Welch et al., 2003; Yang et al., 2004a; Fayard et al., 2005), with interactions that are indirect, or remain to be analyzed in detail, as dashed arrows (Le Blanc et al., 2004; Viard et al., 2004; Brady et al., 2006; Heo et al., 2006; Sun et al., 2006). Biological processes controlled in a PIP3dependent manner are shown in bold, positioned close to their implicated regulators

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cell growth, division, survival, and differentiation to produce highly polarized cells, whilst in postmitotic cells, sensitivity to neurotransmitters and other stimuli, protection from stress and the neuronal plasticity that underpins learning and memory are all influenced by the activity state of PI3K signaling pathways.

1.2 Class I PI3K Signaling is Antagonized by PTEN PIP3 is metabolized by two classes of enzymes: 5-phosphatases, which function to generate a second signal molecule with a distinct pattern of binding partners (see earlier); and 3-phosphatases which inactivate the signal by regenerating its abundant precursor, PI(4,5)P2. PTEN is the only PIP3 3-phosphatase known to date that is widely distributed amongst mammalian tissues and cell types. PTEN was first identified in 1997 as a candidate tumor suppressor in brain, prostate, breast, and kidney tumors (Li et al., 1997; Steck et al., 1997). A wealth of subsequent research has confirmed PTEN as an important tumor suppressor in many cell lineages, and as one of the most commonly mutated genes in human cancers (Leslie and Downes, 2004; Chow and Baker, 2006). However, intense research into the functions of PTEN has now revealed that it is not only an important tumor suppressor, but also a physiological regulator of many diverse biological processes apparently unrelated to tumor development. Strong evidence indicates that PTEN mediates many of its functions through its lipid phosphatase activity, metabolizing PIP3 back to PI(4,5)P2, acting in direct opposition to PI3K. As mentioned, PI3K and PIP3 have many recognized functions, especially in the regulation of cell survival, proliferation, and growth, but also controlling a wider range of cellular functions that includes many cytoskeletal rearrangements and cell migration, the generation of reactive oxygen species (ROS) by NADPH oxidases, insulin stimulated glucose uptake, some forms of phagocytosis etc. Thus, most, and possibly all of these functions are in turn regulated by PTEN. In addition, PTEN has several other potential mechanisms of action, including protein phosphatase activity and nonenzymatic roles, and there is strong evidence that some of its effects are mediated by these latter mechanisms independently of PIP3 (Myers et al., 1997; Gildea et al., 2004; Raftopoulou et al., 2004; Ji et al., 2006; Leslie et al., 2007; Shen et al., 2007). On the other hand, the physiological and pathological significance of these PIP3-independent roles for PTEN is currently unclear.

1.3 The PTEN Protein The PTEN protein contains 403 amino acids, comprising an N-terminal phosphatase domain related to the protein tyrosine phosphatase family, a more C-terminal calcium independent lipid-binding C2 domain, and an unstructured C-terminal tail of approximately 50 amino acids. This C-terminal tail contains several phosphorylation sites and a protein-binding site at the extreme C-terminus, which mediates interactions with a subset of PDZ domain containing proteins. These binding partners include the MAGI proteins 1,2, and 3 (Wu et al., 2000a, b) dPAR-3 (bazooka) (von Stein et al., 2005; Pinal et al., 2006) and possibly Discs large (Adey et al., 2000), all of which besides MAGI-3 have been shown to localize to synapses and may also play roles in the determination of cellular polarity (Muller et al., 1995; Strochlic et al., 2001; Nishimura et al., 2002; Ruiz-Canada et al., 2004; Sumita et al., 2007). PTEN has phosphatase activity in vitro against both phosphoinositides and protein substrates, but the in vivo significance of PTEN action on substrates other than PIP3 remains to be demonstrated. PTEN appears to be a soluble protein with a significant cytosolic pool that displays regulated localization to the plasma membrane, nucleus and specific protein complexes, such as adherens junctions (Das et al., 2003; Gil et al., 2006; Pinal et al., 2006; Vazquez et al., 2006; Leslie et al., 2007; Trotman et al., 2007). In many cases this control over protein localization is linked to the regulatory modification of the protein that occurs via such mechanisms as phosphorylation, ubiquitination and oxidation (Lee et al., 2002; Leslie et al., 2003; Vazquez et al., 2006; Wang et al., 2007). The regulation of PTEN function is rather poorly understood, especially in the brain, although recent work has shown that its activity can be inhibited in neurons and neuronal cell lines by phosphorylation of the C-terminal tail induced by stimulation with nerve growth factor (NGF) or leptin (Arevalo and RodriguezTebar, 2006; Ning et al., 2006).

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PI3K/PTEN Signaling Controlling Neuronal Morphogenesis

During embryonic and early postnatal development, PI3K signaling is vital for the establishment of neuronal morphologies, playing important functions during neuronal polarization, neurite outgrowth, branch complexity, and synapse formation. Similarly, once neuronal circuits are formed, PI3K signaling is essential to preserve the morphology of individual neurons in a circuit.

2.1 Breaking Symmetry – PI3K/PTEN Signaling During Neuronal Polarization After terminal differentiation of a precursor cell, neuronal processes are formed that subsequently segregate into morphologically and functionally discrete cellular domains, the axons and dendrites. The involvement of PI3K and PIP3 in the determination of neuronal polarity first emerged in the best-characterized cellular models used for studying neuronal morphogenesis, the in vitro culture of hippocampal neurons (Dotti et al., 1988) (> Figure 13-3). Shortly after plating, hippocampal neurons produce several indistinguishable short neurite processes (stage 2), which remain quiescent until one neurite rapidly elongates and adopts axonal characteristics (stage 3). Only then the remaining neurites elaborate and aquire dendritic characteristics (stage 4) and will eventually, after approximately 7 days in vitro culture, establish synaptic contacts (stage 5) (Reviewed in Craig and Banker, 1994). During the initial stage of polarization, PI3K has been shown to be selectively activated within the growth cone at the tip of the future axon (Shi et al., 2003; Menager et al., 2004) (> Figure 13-3). Rac1 and Cdc42 appear to participate in a positive feedback loop with PI3K and PIP3, establishing a defined axonal region with high PIP3 and high levels of active Rac1-GTP (Da Silva et al., 2005). Spatially restricted accumulation of PIP3 recruits a polarity protein complex consisting of mPar3/Par6 and aPKC (Shi et al., 2003). Both application of PI3K inhibitors and overexpression of PTEN fail to localize the complex and have been shown to disrupt neuronal polarization (Shi et al., 2003; Menager et al., 2004; Jiang et al., 2005). In vitro, Neurons will polarize and exhibit localized enrichment of PIP3 in the absence of extracellular factors (Menager et al., 2004; Arimura and Kaibuchi, 2007). In addition to spontaneous restricted signaling to the future axon, PI3K activity downstream of IGF-1R activation has recently been shown to drive axonal specification, as inhibition of IGF-1R by siRNA knockdown results in a failure to specify an axon, and this could not be rescued by stimulation with BDNF or NGF (Sosa et al., 2006). In addition, extracellular exposure of a single neurite to laminin promotes neurite growth and favors axon formation involving PI3K signaling (Lein et al., 1992; Esch et al., 1999; Menager et al., 2004). Thus, localized activation of IGF-1R and/or laminin appears to drive PI3K mediated initiation of elongation and specification of the axonal process. Establishing neuronal polarity in hippocampal neurons has been suggested to require both the protein kinase, GSK3b, and its substrate CRMP-2 (Arimura et al., 2004; Cole et al., 2004; Yoshimura et al., 2005) and it has been known for many years that GSK3 can be regulated by N-terminal PI3K dependent Akt-mediated phosphorylation (Cross et al., 1995). Despite this, GSK3 phosphorylates many of its substrates in a PI3K and Akt independent manner and the role of PI3K and Akt-dependent GSK3 regulation in the determination of neuronal polarity is currently controversial. Mice in which the Akt phosphorylation sites in GSK3a and b are mutated to alanines lack detectable regulation of either GSK3 kinase by PI3K and Akt, develop normally, and the establishment of polarity in cultured hippocampal neurons from these mice, although GSK3-dependent, appears normal, arguing against a widespread obligate role for the Akt-dependent regulation of GSK3 in axonal differentiation (McManus et al., 2005; Gartner et al., 2006).

2.2 Spatially Restricted PIP3 Accumulations Regulate Cell Polarity in Nonneuronal Cells The morphological polarization of neurons has crucial functional implications that are essential for the function of neuronal networks. Evidence from other model systems, such as the study of neutrophil and

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. Figure 13-3 The roles of PI3K/PTEN signaling during polarization and development of hippocampal neurons. A schematic diagram is shown of stages 2–5 of the normal morphological neuronal development of a rat hippocampal neuron (lower panel) and the proposed experimentally defined roles of PI3K/PTEN signaling (upper panel). Overexpression of PTEN inhibits axon formation (Shi et al., 2003; Jiang et al., 2005). Downregulation of PTEN by siRNA (Jiang et al., 2005) or overactivation of PI3K by expression of myr-p110 PI3K (Yoshimura et al., 2006) increases the number of neurons with multiple axons. The growth factors BDNF and NT-3 increase the number of axonal branches in stage 3 neurons by activation of PI3K signaling (Yoshimura et al., 2005), whilst NGF increases axonal length by inhibition of PTEN through C-terminal phosphorylation (Arevalo and Rodriguez-Tebar, 2006). Dendritic branch morphology is greatly increased by Reelin or BDNF through activation of PI3K (Jaworski et al., 2005; Kumar et al., 2005; Jossin and Goffinet, 2007). In the absence of growth factor stimulation overactivation of PI3K signaling by expression of myr-p110 PI3K or siRNA knockdown of PTEN increases dendritic branch complexity and appearance of dendritic filopodia (Jaworski et al., 2005; Kumar et al., 2005). On the contrary, pharmacological inhibition or expression of dominant-negative p85 PI3K reduces the number of dendritc protrusions (Kumar et al., 2005). Neurites, axons, and growth cones are labeled, and PIP3 enriched membrane domains are shown in light gray

dictyostelium chemotaxis, has shown that PI3K is also a key mediator during cell polarization and directed motility in a number of cellular systems (Funamoto et al., 2002; Wang et al., 2002). Analyses of chemotaxis, first in the slime mould Dictyostelium, and later in mammalian neutrophils and fibroblasts showed that PIP3 is enriched at the leading edge of these polarized cells (Parent et al., 1998; Meili et al., 1999; Haugh et al., 2000; Servant et al., 2000), and PTEN has been found to be enriched at the back of chemotaxing dictyostelium cells and also probably neutrophils (Funamoto et al., 2002; Iijima and Devreotes, 2002; Li et al., 2003b; Li et al., 2005). It is thought that a positive feedback loop involving PIP3 accumulation is sufficient to activate the polarization process and motility even in the absence of chemoattractant (Niggli, 2000; Weiner et al., 2002), which has been suggested to depend additionally on rho-GTPases and possibly F-actin (Wang et al., 2002; Weiner et al., 2002). Other potentially significant mechanisms in this regard are the apparent roles for PIP3 in the activation of PLCg1 and the inhibition of RhoA (Bae et al., 1998; Falasca et al., 1998; Krugmann et al., 2004; Krugmann et al., 2006). Work in diverse systems, including polarized

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epithelial cells (Martin-Belmonte et al., 2007), cells undergoing division (Janetopoulos et al., 2005), and polarizing neurons (Shi et al., 2003) has now established a role for PI3K, PIP3, and PTEN in cellular polarization. However, although parallels can be drawn between these diverse systems, the interplay between PI3K signaling and other signaling systems that establish and regulate cellular polarity remains to be clarified, and may differ between cell types (Comer and Parent, 2007). Significantly, once polarity is established in neurons and epithelial cells, it appears to be retained in the absence of applied external polarizing signals, in contrast to dictyostelium cells or neutrophils.

2.3 PI3K/PTEN Signaling During Extension and Navigation of Neurites Following the establishment of neuronal polarity, extension of axonal and dendritic processes are critical for the establishment of appropriate connectivity. A body of work underscores the significance of growth factor signaling in the promotion of neurite outgrowth by activation of PI3K signaling. In PC12 cells, local accumulation of PIP3 and subsequent activations of Rac1/Cdc42 play a critical role in NGF-induced neuritogenesis and neurite outgrowth (Aoki et al., 2004; Aoki et al., 2005; Nakamura et al., 2005), a signaling module similar to the localized positive feed back loop that operates during the establishment of neuronal polarity (see earlier). Localized and prolonged activations of PI3K signaling necessitate desensitization of this signaling pathway by a negative regulator. This is achieved, at least during neurite outgrowth responses in PC12 cells, by inhibition of the NGF–PIP3–Rac1/Cdc42 signaling network through SHIP2 and PTEN (Aoki et al., 2007). Thus, increased turnover of PIP3 appears to maintain a growth cone in a ‘‘motile state,’’ leading to enhanced outgrowth responses, an idea that is supported by the observation that acute inhibition of PI3K indeed antagonizes neurite outgrowth and induces a growth cone collapse response (Sanchez et al., 2001; Atwal et al., 2003; Dijkhuizen and Ghosh, 2005; Chadborn et al., 2006). A fundamental feature of the outgrowth of axonal processes is their ability to follow stereotypic paths to appropriate targets. Axonal guidance is controlled by interactions between the growth cone and guidance molecules present in the local microenvironment, several of which have been shown to evoke directional responses dependent upon of PI3K signaling. In an early study by Ming et al., Netrin-mediated attractive turning in the pipette assay was desensitized in the presence of pharmacological inhibitors for PI3K (Ming et al., 1999), suggesting a dependence of PI3K activation during Netrin function. This in vitro work is best supported by work in C. elegans, where UNC-6/Netrin induces neuronal asymmetry and defines the site of axon formation through localized accumulation of the PI(3,4)P2 binding protein lamellipodin. Lamellipodin membrane accumulation is driven by both PI3K and PTEN (Adler et al., 2006; Chang et al., 2006). On the contrary, sustained activation of PI3K has been shown to reverse Sema3F or Sema3A induced growth cone collapse (Atwal et al., 2003), and we have recently shown that Sema3A suppresses PI3K signaling in a PTEN-dependent fashion (Chadborn et al., 2006). Thus, axonal growth stimulators (such as growth factors or Netrin) and inhibitors (such as class III Semaphorins) converge to modulate levels of membranous PIP3 and define the fine-tuning between axonal growth and inhibition.

2.4 Neurite Branching and Dendritic Morphogenesis are Controlled by PI3K/PTEN Signaling The proper functioning of the nervous system relies not only on the appropriate navigation of neurites, but also on their complex branching and arborization behavior. Both dendrites and axons are usually branched, with the extent of branching customized to the functional needs of the neural networks they assemble. Several ligand/receptor systems regulate axonal and/or dendritic branching in different neuronal populations. For example, NGF and BDNF control neurite branching in sympathetic neurons and retinal ganglion cells, respectively, whereas neurotrophin-3 (NT-3) controls terminal branching complexity in DRG axons (Hoyle et al., 1993; Cohen-Cory and Fraser, 1995; Lentz et al., 1999). NT-3 also enhances axonal branching in hippocampal neurons, whilst BDNF evokes increases in branch complexity of axonal and dendritic processes in the same cell type. Conversely, factors with inhibitory effects on neurite growth and guidance

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have been shown to destabilize already existing neurite branches. Sema3F, an inhibitory axon guidance molecule, is implicated in the pruning of branches during hippocampal development (Bagri et al., 2003). Despite its importance, our understanding of the molecular mechanisms controlling neurite branching is limited; however, several studies have identified an involvement of PI3K in signaling by many of the aforementioned branch-controlling factors (Jaworski et al., 2005; Kumar et al., 2005; Brandt et al., 2007; Jossin and Goffinet, 2007). Importantly, direct manipulation of the activity of PI3K or levels of PTEN changes the extent of neurite branching, indicating that levels of PIP3 are sufficient to determine branch complexity of neuronal processes. For example, in hippocampal neurons cultured in vitro, PI3K signaling plays a pivotal role in the regulation of many aspects of dendrite formation. Chronic inhibition of PI3K reduces dendrite size and dendritic complexity, as well as density of dendritic filopodia and spines (Kumar et al., 2005). Constitutively active mutants of Ras, PI3K, and Akt, or loss of PTEN by siRNA induces growth and elaboration of dendrites (Jaworski et al., 2005; Kumar et al., 2005; Jaworski and Sheng, 2006). Downstream effectors of PI3K/PTEN activity in this process include the Akt/mTOR/p70S6K pathway regulating protein translation (Jaworski et al., 2005; Kumar et al., 2005).

3

PI3K/PTEN Signaling in Normal Brain Function and Disease

The central significance of PI3K/PTEN to normal brain function is underscored (Gimm et al., 2000), (Li et al., 2003a; Greer and Wynshaw-Boris, 2006; Kim and Mak, 2006), by the high incidence of neurological deficits that occur as a result of mutations in PTEN or in components downstream of PI3K signaling. For example, PTEN mutations occur in four human syndromes, which result in macrocephaly associated with symptoms of autistic spectrum disorders (ASD) and mental retardation (Goffin et al., 2001; Eng, 2003). Individuals carrying inactivating mutations of PTEN develop Cowden disease and the propensity to develop tumors can coincide with neurological defects such as mental retardation, ataxia, and seizures (Liaw et al., 1997). Given that loss of PTEN in vitro leads to distinct and lasting effects on neurite outgrowth and synaptogenesis, it is not surprising that inactivating mutations in PTEN in humans are likely to affect normal neuronal circuit formation and consequentially affect cognitive functions.

3.1 Learning and Memory The evidence that PI3K/PTEN signaling is required for the maintenance of the synaptic plasticity that underpins current concepts of learning and memory stems mainly from the use of pharmacological inhibitors of PI3Ks. In many areas of the brain, including the hippocampus, two opposing forms of synaptic plasticity, long term depression and long term potentiation (LTD and LTP, respectively), have been defined, can be induced following different conditioning protocols applied to postsynaptic neurones and require Ca2+ influx through excitatory NMDA receptors. Interestingly both LTD and LTP are facilitated by insulin and blocked by PI3K inhibitors. Although the mechanism is not understood, it has been proposed that the effects of insulin might involve enhanced expression of PSD-95, a protein that functions as a scaffold linking the NMDA receptor with additional signaling components (Sheng, 2001). Further pharmacological studies in rats have linked PI3K signaling to the development of fear conditioning and memory retrieval and extinction (Lin et al., 2001; Chen et al., 2005; van der Heide et al., 2006), but as noted below, such studies must be viewed with caution given the current understanding of the off target effects of the inhibitors used in these experiments.

3.2 Seizure and Epilepsy Several lines of evidence link PI3K signaling in the nervous system to seizures and epilepsy. For example, the antiepileptic drug, carbamazepine was found to enhance the activity of the type 3 glutamate transporter via

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a PI3K-dependent process that could be inhibited by wortmannin and LY294002 (Lee et al., 2005) and kainate-induced seizures are exacerbated by such PI3K inhibitors (Henshall et al., 2002). These observations, however, suffer from limitations related to potential off-target effects (both wortmannin and LY294002, for example, inhibit the protein kinase, casein kinase 2, Class III, and some Class II PI3Ks as well as Class I PI3Ks, while LY29002 is a fairly potent inhibitor of mTOR) and, for the time being, must be considered with caution. Moreover, since kainate-induced seizures result from engagement of the Bcl-2 associated death protein (BAD) pathway leading to apoptotic cell death and PIP3 signaling antagonizes this pathway, this outcome could have been predicted and may be specific to the mechanism of seizure induction. More compelling evidence comes from two independent studies of mice in which PTEN was specifically deleted from the brain using the Cre-loxP system (Backman et al., 2001; Kwon et al., 2001). These mice have a phenotype that is strikingly similar to Lhermitte–Duclos disease associated with Cowden disease patients and exhibit, amongst other features, seizures and ataxia associated with cerebellar lesions that eventually cause them to die in 29 weeks. These features appear to result from defects of growth and migration of granule cells rather than being neoplastic in nature.

3.3 PTEN/PI3K Signaling and Autism Spectrum Disorders Clinically highly relevant neuronal functions of PI3K/PTEN are related to its association with ASD. ASD is a genetically complex neurodevelopmental syndrome characterized by abnormalities in language development and social interaction, that coincides with restricted stereotypic and repetitive patterns of behaviors (McCaffery and Deutsch, 2005; Persico and Bourgeron, 2006). A subset of individuals with ASD show a substantially accelerated brain growth during a critical postnatal time window, leading to a statistical overrepresentation of individuals with enlarged brain volumes (macrocephaly), which may, however, disappear at later developmental stages (Fombonne et al., 1999; Courchesne and Pierce, 2005; McCaffery and Deutsch, 2005). The disturbance of early brain growth coincides with a time of critical events in neuronal circuit formation. Indeed, the elaboration of neurite processes, and synaptic connectivity and the extent of myelination has been shown in different studies to be affected in individuals with ASD (Herbert et al., 2004; Herbert, 2005). Thus brain overgrowth during a restricted postnatal period is at present the most replicated pathology of ASD (Herbert, 2005). Deregulation of PI3K/PTEN signaling is currently thought of as a contributing factor associated with ASD, partially because of its role in regulation of neuronal morphogenesis, synapse formation, and synapse plasticity. Indeed, a recent study focussing specifically on subjects with autism that exhibited severe macrocephaly found 23% of cases with germline mutations of PTEN (Butler et al., 2005). However, caution has to be taken as in patients carrying germline mutations of the PTEN gene in Cowden disease, ASD was never diagnosed (Liaw et al., 1997). More convincingly, perhaps, tuberous sclerosis, resulting from mutation of the tuberous sclerosis complex (TSC) 2 protein, a target for phosphorylation by Akt, is associated with a remarkable 25–50% incidence of autism (Wiznitzer, 2004). The TSC1/2 complex links PI3K signaling to mTOR, which functions to coordinate cell growth and division with nutritional status (see > Figure 13-1) and may explain the apparent link between brain size in children and ASD. The idea that accelerated brain growth as a consequence of abberant PI3K/PTEN signaling may underlie postnatal neurodevelopmental disorders including ASD is further supported by different conditional PTEN-deficient mice. Homozygous deletions of PTEN in mice were quickly found to result in early embryonic lethality (Di Cristofano et al., 1998; Suzuki et al., 1998), and thus numerous mouse lines were engineered in which PTEN-loss is achieved in a tissue-specific and temporal manner under the control of different promoters (Backman et al., 2001; Groszer et al., 2001; Kwon et al., 2001; Marino et al., 2002; Fraser et al., 2004; Yue et al., 2005; Kwon et al., 2006). The general outcome shows that genetic inactivation of PTEN leads to cellular hypertrophy, and the consequential increase of tissue sizes in areas of PTEN deficiencies. In one of the most compelling studies PTEN deficiency was restricted to a subpopulation of fully differentiated neurons that had already established synaptic contacts within the cerebral cortex and hippocampus. This late loss of PTEN induced dramatic changes in neuronal circuitries, generating ectopic dendrites and axonal tracts, and increased the number of synapses. The appearance of enlarged neuronal

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soma and progressive macrocephaly, both of which are reminiscent of ASD in humans (Kwon et al., 2006), together with the development of neurological symptoms that include deficits in social interactions, seizures, and decreased learning suggest that these mice may directly provide a causal link between PTEN deficiency and ASD (Kwon et al., 2006). However, one has to take into consideration that autism is thought to result from developmental defects, whilst the mouse conditional PTEN knockout phenotype results from postmitotic effects in the targeted neurons. Given the previous work linking PI3K signaling via TSC1/2 and mTOR to autistic endpoints, it would be interesting to examine effects of centrally administered rapamycin (which inhibits the activity of mTOR complex 1) on the development of these features using the mouse model.

3.4 PTEN is Required for Addictive Responses to Drugs of Abuse A key role for PTEN, distinct from that of antagonizing PI3K signaling, is emerging in the dephosphorylation of regulatory sites on cytosolic domains of cell surface receptors. As PTEN is targeted to membranes in order to efficiently dephosphorylate PIP3, its ability to regulate protein substrates seems to depend on specific protein/protein interactions. Recent evidence suggests that PTEN interacts directly with the 5-HT2C receptor in the ventral tegmental area (VTA) of the brain via a short peptide sequence within the receptor’s third intracellular loop. This receptor normally acts to repress the firing of dopaminergic neurons in the VTA that is triggered by nearly all addictive drugs of abuse including cocaine, alcohol, and the active ingredient of marijuana. By generating a cell-penetrant peptide to mimic this sequence, Ji and colleagues were able to disrupt the interaction of the receptor with PTEN in PC12 cells and in rats after injection of the peptide into the VTA (Ji et al., 2006). The results of these experiments suggest that PTEN specifically targets a site in the 5-HT2C receptor that becomes phosphorylated upon agonist stimulation, an event that appears to be required to maintain the receptor in an activated state. Hence PTEN normally acts to desensitize 5-HT2C receptor function and is permissive for the rewarding effects of drugs of abuse, and led the authors to suggest that specifically disrupting the interaction of PTEN with 5-HT2CR would be a good strategy to combat addiction.

4

PI3K/PTEN Signaling in the Central Control of Metabolism

Lipid phosphatases, such as PTEN and SHIP2, by their substrate specificity in hydrolyzing PIP3 and ability to act as negative regulators of PI3K signaling were expected to be key regulators of the PI3K-dependent processes underlying the actions of insulin on glucose homeostasis. Indeed, impaired insulin-stimulated PI3K-PKB signaling is central to the development of insulin resistance in diabetes (Zdychova and Komers, 2005), and downregulation of PTEN in various peripheral tissues improves glucose disposal and prevents diabetes (Sasaoka et al., 2006; Sutherland and Ashford, 2008). However, little information is available at present on the effects of PTEN manipulation on central control of glucose or energy homeostasis. Leptin and insulin act in the brain to reduce food intake and body weight and utilize PI3K signaling pathways, amongst others, to elicit anorexigenic outputs in hypothalamic neurons and circuits (Niswender et al., 2004). Consequently, it is predicted that manipulations which enhance PI3K signaling should lead to decreased food intake and negative energy balance. In support of this concept, SHIP2 knock-out mice are protected from obesity when challenged by a high fat diet (Sleeman et al., 2005), although it has not yet been determined whether this effect is mediated in the CNS. One of the major sites for leptin and insulin action, in relation to energy homeostasis, is the proopiomelanocortin (POMC)-expressing neurons of the hypothalamic arcuate nucleus, a major anorexigenic mediator (Cone, 2005). In these neurons leptin and insulin signaling converges with activation of the PI3K pathway (Xu et al., 2005) and POMC neuron specific reduction in PTEN resulted in mice displaying hyperphagia and for female mice, diet-sensitive obesity and leptin resistance (Plum et al., 2006). Although this result appears counter-intuitive to that expected for enhancement of PI3K signaling pathways, electrophysiological analysis of these PTEN deficient POMC neurons showed that they were electrically

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silenced, thus reducing anorexigenic signaling. While it may be argued that both leptin and insulin signals through the PI3K pathway are negatively regulated by PTEN, the relation between these hormones and regulation of cellular PIP3 levels appears to be more complex. It has been shown that leptin, but not insulin, can increase the phosphorylation of PTEN and inhibit its lipid and protein phosphatase activity, thus increasing PIP3-dependent signaling processes independently from stimulation of PI3K (Ning et al., 2006). Thus, further analysis of the role of PTEN in leptin and insulin signaling pathways may provide important and novel insights in relation to not only obesity and diabetes, but also pathologies involving aberrant cell survival and death signals.

5

PI3K/PTEN Signaling and Neurodegenerative Disease

5.1 Ischemic Brain Injury/Stroke Hypoxic-ischemic brain injury underlies the damage induced by stroke, the third most common cause of death in developed countries. Ischemic neuronal death is thought to be due to a combination of excitotoxicity, oxidative stress, and apoptosis (Won et al., 2002; Zheng et al., 2003). Evidence from the main experimental model, transient focal/global ischemia followed by reperfusion in rodents, indicates an important role for involvement of cell survival/death signaling pathways (Won et al., 2002; Saito et al., 2005). Mitogen-activated protein kinases such as p38 and JNK are activated in response to cerebral ischemia and may mediate neuronal cell death (Won et al., 2002; Irving and Bamford, 2002; Saito et al., 2005). Additionally, activation of the PI3K-Akt pathway has been shown to be neuroprotective following ischemic insult (Wick et al., 2002; Fukunaga and Kawano, 2003; Endo et al., 2006), and can act to negatively regulate JNK signaling and JNK-mediated apoptosis (Yang et al., 2004b; Hui et al., 2005). Thus, alterations in the level or activity status of PTEN by an ischemic insult could influence the balance between these signaling systems and the survival outcome of the neuron. Using the focal or global ischemic-reperfusion model, various studies have demonstrated that the phosphorylation of PTEN is increased in the hours following ischemic insult, consistent with inactivation of the protein (Omori et al., 2002; Choi et al., 2004; Choi et al., 2005). Omori et al. (2002) also demonstrated a significant reduction in the overall levels of PTEN protein in the ischemic core (Omori et al., 2002). Various studies have suggested that the down regulation of PTEN is associated with a neuroprotective role (Gary and Mattson, 2002; Lee et al., 2004; Zhu et al., 2006). In hippocampal neurons, reduced PTEN expression was associated with increased Akt activity and decreased neuronal death following transient global ischemia (Ning et al., 2004). Furthermore, pharmacological inhibition of PTEN has been reported to be associated with increased Akt activity and reduced neuronal damage (Lee et al., 2004; Wu et al., 2006). An intriguing and perhaps surprising additional role for PTEN has been suggested in relation to ischemic injury. This involves the possibility that, rather than being inhibited by oxidation, mitochondrially located PTEN actually participates in ischemia-reperfusion-induced production of ROS. Using rat cultured hippocampal neurons and the human dopamine containing cell line SH-SY5Y, Zhu et al (2006, 2007) demonstrated that antisense or siRNA knockdown of PTEN reduced ROS generation and provided neuroprotection against both ischemic and neurotoxin challenges (Zhu et al., 2006; Zhu et al., 2007). This may provide an intriguing link between mitochondrially located PTEN and neuronal injuries associated with stroke and Parkinson’s disease. These data indicate that reduced PTEN expression and/or function and consequent enhancement of PI3K-Akt signaling may act as a cellular protective mechanism in response to injury. However, recent work suggests that the reduction in PTEN function associated with ischemia may be temporary. Examining ischemic-reperfusion injury in the CA1 region of the hippocampus, demonstrated that in the first few hours following the ischemic insult phosphoserine PTEN levels decreased concomitant with increased phosphotyrosine PTEN, which resulted overall in decreased phosphatase activity and increased degradation of PTEN (Zhang et al., 2007b). This produced a transient increase in Akt and decrease in JNK signaling, but was followed by a delayed increase in PTEN levels with consequent reduced Akt and increased JNK signaling. Interestingly, inhibition of protein tyrosine phosphatases, which increased tyrosine

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phosphorylation of PTEN, or antisense knockdown of PTEN resulted in a longer lasting reduction of PTEN protein levels. This not only enhanced PKB signaling and reduced JNK signaling (Zhang et al., 2006a), but also exerted a neuroprotective outcome with reduced levels of neuronal death (Wu et al., 2006; Zhang et al., 2007b). Therefore, it may be argued that acute PTEN inhibition, with consequent upregulation of the prosurvival pathway PI3K-Akt in concert with downregulation of cell death pathways such as JNK, could contribute to improved neuronal survival following ischemia-reperfusion injury. However, the results from Zhang et al. (2007b) indicate that the downregulation of PTEN is transient, lasting less than 1–2 days, and that recovery of PTEN protein levels in the days following the ischemic insult is still capable of causing neuronal death. Consequently, a long-term inhibition of PTEN may be beneficial to protect against delayed neuronal damage initiated by the transient ischemic insult. In relation to this, ischemic preconditioning, a powerful endogenous protection mechanism against ischemic-reperfusion injury that involves increased signaling through the PI3K-Akt pathway, has been shown to result in depression of PTEN protein levels in rat isolated heart (Cai and Semenza, 2005). Although this study has been criticized with respect to the short duration of the ischemic insult, this is clearly an important issue and requires further studies on both cardiac muscle and neurons. Such a chronic inhibition of PTEN is not without its dangers of course, as prolonged loss of PTEN levels or function may have significant negative consequences for the neuron (e.g., cell hypertrophy, inappropriate axonal outgrowth) or in the case of glia, inappropriate cell proliferation. In addition, a long-lasting decrease of PTEN function, in concert with upregulated PI3K-PKB signaling, may also increase the risk in susceptible individuals to other neurodegenerative conditions such as Alzheimer’s disease. These studies show the importance of PTEN’s lipid phosphatase activity in neuroprotection. However, work using an oxygen glucose starvation model of ischemic injury, indicates that PTEN may act in neuroprotection in a dual specific manner, involving both its lipid and protein phosphatase activity. PTEN has been shown to associate directly with the NMDA receptor subunits NR1 and NR2B at extrasynaptic sites in hippocampal neurons and has been suggested to regulate NMDA receptor function (Ning et al., 2004). Whilst the selective loss of the lipid phosphatase activity alone increased Akt and BAD phosphorylation, the neuroprotective effects of PTEN-loss were enhanced by diminishing extrasynaptic NMDA receptor activity, which required the additional lack of the protein phosphatase activity (Ning et al., 2004). Thus, PTEN’s involvement in controlling neuroprotection seems twofold, however, whether PTEN’s phosphatase activity directly targets NMDA receptors for dephosphorylation remains to be tested.

5.2 Alzheimer’s Disease Alzheimer’s disease (AD) is a progressive neurodegenerative disorder and the most prevalent form of dementia, which affects nearly 2% of the population of industrialized countries (Blennow et al., 2006). Characteristic features of the brain in patients having AD are the presence of excessive numbers of ‘‘plaques’’ and ‘‘tangles.’’ The extracellular plaques are due to the abnormal accumulation of amyloid b-peptide (Ab) a 39–42 amino acid peptide generated by sequential proteolysis of amyloid precursor protein (APP), with cleavage by the beta site APP-cleaving enzyme (BACE1), the rate-limiting step. The intracellular neurofibrillary tangles (NFTs) are aggregates of the microtubule-associated protein tau, which has undergone abnormal levels of phosphorylation. Several causative mutations have been found linking specific genes with AD, but the vast majority of cases of AD are sporadic, with certain behavioral, dietary, and environmental factors (e.g., high calorie, high fat diet, sedentary lifestyle) increasing the risk of AD (Mattson, 2004). Clearly, the pathophysiological events that cause the increased levels of Ab and hyperphosphorylated tau are likely to be key to elucidating the mechanisms underlying the neuronal cell death associated with AD and related neuropathologies. Recent studies indicate that the PI3K-Akt-GSK3 pathway plays a pivotal role in the regulation of both Ab production and tau phosphorylation, and may provide a link between these seemingly disparate pathologies and neuronal cell loss. Although early reports suggested the possibility of impaired PI3K-Akt signaling in the brain of patients having AD (Hanger et al., 1992; Mandelkow et al., 1992; Lovestone et al.,

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1994; Brownlees et al., 1997; Sun et al., 2002; Phiel et al., 2003; Ryder et al., 2003), recent studies of human postmortem brain report increased levels of phosphorylated Akt associated with the disease (Pei et al., 2003; Rickle et al., 2004). Additionally, PI3K-Akt activation has been reported to protect against Ab-induced neurotoxicity in cells (Martin et al., 2001; Wei et al., 2002) and is associated with reduced levels of amyloid plaque formation in a mouse model of AD (Stein and Johnson, 2002). It is therefore plausible that the observed increased activity of the PI3K-Akt pathway is an initial survival response to neuronal injury which has ultimately deleterious consequences for certain mature neurons, promoting outcomes that result in AD pathology and eventual neurodegeneration. In line with this possibility, studies in postmortem brain (Griffin et al., 2005), cell lines (Kerr et al., 2006; Zhang et al., 2006b; Zhang et al., 2006c), and primary cultured neurons (Zhang et al., 2006b) have established the relationship between PTEN-loss, enhanced PI3K/Akt responsiveness, and Tau phosphorylation. Indeed, Akt activation appears to increase phosphorylation of tau selectively at Ser214 (Ksiezak-Reding et al., 2003; Kyoung Pyo et al., 2004; Griffin et al., 2005), which along with the neighboring Thr212 site forms the AT100 epitope, considered to be a specific marker for AD (Matsuo et al., 1994) and overexpression of wild type PTEN reduces levels of tau phosphorylation, increases tau–microtubule association, and decreases formation of tau aggregates (Kerr et al., 2006; Zhang et al., 2006b). Similar results were observed using a mutant form of human tau (tau FTDP-17 mutation), which is closely associated with dementia, where interference with PTEN function induced the mutant tau to form aggregates (Zhang et al., 2006b; Zhang et al., 2006c) and resulted in impaired neurite outgrowth (Zhang et al., 2006b). Although these latter reports have mainly utilized recombinant proteins and cultured cell systems, a limited study of three cases of AD patients’ brains showed significantly reduced levels of PTEN associated with Ser214 phosphorylated tau (Zhang et al., 2006b). Moreover, using PTEN-conditional knockout mice, ablation of PTEN in embryonic or mature cerebellar Purkinje cells resulted in increased Akt levels, tau hyperphosphorylation (at multiple sites, including Ser214), and increased cytoplasmic deposition of fibrillary inclusions, culminating in the chronic progressive loss of Purkinje cells in an age dependent manner (Marino et al., 2002; Nayeem et al., 2007). Interestingly, PTEN may also provide a link to encompass Ab pathology. The production of Ab following cleavage of APP by BACE1 requires a secondary cleavage by the g-secretase complex, which consists of at least four protein components. Of these, the presenilins (PS1 and PS2) play an important role in the production of Ab and progression to the AD phenotype. PS mutations account for the vast majority of familial AD cases, and many result in increased levels of Ab (Shen and Kelleher, 2007). Indeed, unlike mouse models that overexpress mutant human APP, conditional knockout of PSs in adult mouse cerebral cortex closely recapitulates the dementia and neurodegeneration seen in human AD (Shen and Kelleher, 2007). Interestingly, a recent report shows that PS1 and PS2 knockout in mouse embryonic fibroblasts and mouse forebrain reduces the level of PTEN, whereas restoration of PS into deficient cells, or PS overexpression, increased PTEN, actions apparently independent of g-secretase activity (Zhang et al., 2008). Thus, the question of whether changes in PTEN levels and activity connect aberrant neuronal signaling with AD linked pathologies of APP mis-processing and tau hyperphosphorylation and aggregation require further investigation.

5.3 Parkinson’s Disease Parkinson’s disease (PD) is the second most prevalent neurodegenerative disorder in industrialized countries with increasing age as the main risk factor. It is characterized by the selective and progressive loss of dopamine containing neurons in the substantia nigra pars compacta, resulting in dysregulated function of motor circuits in the basal ganglia (Savitt et al., 2006). Although there is a genetic component to PD, the vast majority of cases (>90%) are idiopathic. Nevertheless, investigation of the genetic mutations associated with PD, in addition to neurotoxin (e.g., 1-methyl-4phenylpyridnium (MPP+)) induced PD, have been critical to our understanding of the mechanisms underlying this selective neurodegenerative condition (Abeliovich and Flint Beal, 2006; Wood-Kaczmar et al., 2006). Such studies have shown that increased free radical production, oxidative stress, and mitochondrial dysfunction are strongly implicated

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in the pathogenesis of PD (Abou-Sleiman et al., 2006; Savitt et al., 2006). In relation to our theme here, two mutations associated with PD have been demonstrated to have links with PTEN. Firstly, a protein localized to the mitochondrial membrane, PTEN-induced kinase 1 (PINK1) that is linked with familial PD is also associated with mitochondrial dysfunction and cell death (Hatano et al., 2004; Valente et al., 2004). As indicated by its name this protein is transcriptionally activated by PTEN and loss of PINK1 function contributes to PD (Khan et al., 2002; Valente et al., 2004). This connection is supported by studies of PINK1 loss of function mutants in Drosophila, which exhibit severe mitochondrial dysfunction, dopamine neuron degeneration, and increased sensitivity to oxidative stress (Clark et al., 2006; Park et al., 2006). Some clues regarding the likely mechanisms underlying this increased susceptibility to cell death have arisen from investigations on cultured cells. The human neuroblastoma cell line, SH-SY5Y, overexpressing a dominantnegative PINK1 was demonstrated to exhibit a reduced (depolarized) mitochondrial membrane potential and enhanced apoptotic cell death in response to stress (Valente et al., 2004). Conversely, overexpression of wild-type PINK1 was associated with a maintained mitochondrial membrane potential, less activation of caspases, and significantly reduced levels of cell death, effects abrogated in the presence of PD-related mutant PINK1 or catalytically dead PINK1 (Valente et al., 2004; Petit et al., 2005). A second protein, DJ-1, mutations of which are associated with early onset PD (Abou-Sleiman et al., 2003; Bonifati et al., 2003) has a close functional connection to the PI3K-dependent survival pathway, partly through actions on PTEN. In Drosophila, siRNA-mediated reduction of DJ-1 induces neuronal hypersensitivity to oxidative stress (Yang et al., 2005), an outcome shared by DJ-1-deficient mice (Kim et al., 2005b). In these studies loss of DJ-1 and increased cell death were associated with decreased Akt phosphorylation, whereas overexpression of DJ-1 increased Akt phosphorylation (Kim et al., 2005a; Yang et al., 2005). Indeed the neurodegenerative phenotype could be rescued by overexpression of Akt or the PI3K catalytic subunit and enhanced by dominant-negative PI3K or PTEN expression (Yang et al., 2005). It has been suggested that the DJ-1-mediated modulation of the PI3K-Akt survival pathway is due to DJ-1 acting as a negative regulator of PTEN (Kim et al., 2005a). At present it is unclear where the molecular events involving DJ-1, PINK1, PTEN, and PI3K-Akt converge and how these proteins protect neurons from oxidative stress. However, one recent study has provided evidence that DJ-1 and PINK1 may physically associate in the cell, and that PD-associated mutant forms of these proteins display reduced ability to protect against oxidative stress (Tang et al., 2006). DJ-1 may also physically associate with another important PD molecule, parkin, during oxidative stress (Moore et al., 2005) and it appears that all three are physically associated with mitochondria (Valente et al., 2004; Silvestri et al., 2005; Zhang et al., 2005). Accordingly, these proteins, along with PTEN may play important roles in mitochondrial function particularly in relation to the production and handling of reactive oxygen and nitrogen species, and further highlights mitochondrial dysfunction as a pivotal mechanism for PD pathology. Alterations in PI3K-Akt signaling have also been linked to other neurodegenerative disorders, although no role for PTEN has yet been reported. For example, Akt signaling has been associated with two neurological disorders, caused by CAG repeat expansion resulting in the presence of polyglutamine chain containing proteins. In Huntington’s disease (HD), the mutant huntingtin protein, which is responsible for the neurotoxicity of striatal neurons, is phosphorylated by Akt and this process mediates survival of the neurons (Humbert et al., 2002; Rangone et al., 2004). This led to the idea that compromised Akt signaling may be a factor in the progression of HD. In support of this notion, it has been reported that in a rat model of HD the brain levels and activity of Akt are reduced prior to neuron loss, and that Akt may be downregulated in HD patients (Colin et al., 2005). Contrastingly, in spinocerebellar ataxia type 1 (SCA1), Chen et al. (2003) reported that Akt activity drives phosphorylation of the mutant form of ataxin-1 and this results in its association with 14-3-3, leading to ataxin-1 stabilization, accumulation, and eventual neurodegeneration (Chen et al., 2003).

6

Future Perspectives

Current evidence tells us that PI3K/PTEN signaling plays central roles in neurophysiology, and is deregulated, probably causally, in many pathologies of the central nervous system. A detailed understanding of the

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role of PI3K/PTEN signaling in the brain will require significant further experimental advances, especially in the development of in vivo models of the spatially and temporally restricted loss of function of PTEN or PI3K isoforms. Such experiments should also go some way to determining the significance of PTEN’s PIP3-independent mechanisms of action in the brain. Combined with a better grasp of these signaling pathways in terms of how different downstream PIP3 targets regulate different processes, and how the regulation of different PIP3 and PI(3,4)P2 phosphatases impinge on pathway control, this maturation of the field should be reflected not only in a better understanding of the processes in question, but should provide opportunities for the well-founded translation of basic research into therapeutic advances.

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Phosphoinositide-Specific Phospholipase C: Isoforms and Related Molecules

H. Yagisawa

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269

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PLC Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270

3 3.1 3.2 3.2.1 3.2.2 3.2.3

Core Structure of PLC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 Structure of PLCd1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 PLCd Subfamily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 Yeast PLC Regulates Cell Growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 Mammalian PLCd Subfamily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 Gene Knockout of d Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 276

4 4.1 4.2 4.2.1 4.2.2 4.2.3 4.2.4 4.3 4.3.1 4.3.2 4.3.3 4.3.4 4.3.5 4.3.6 4.3.7 4.3.8

Isoform Families that have Auxiliary Domains: Regulation Under the Control of Specific Molecular Interaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 278 PLCg with the SH2/SH3-Containing X-Y Linker Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 278 PLCb Under the Control of Heterotrimeric G Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279 Gbg Dimer Activates PLC Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279 PLCb with C-terminal Extension (tail) Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 280 Expression and Distribution of PLCb Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 282 Gene Knockout Studies of PLCb Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 282 PLCe: Under the Control of Small G Proteins and Heterotrimeric G Proteins . . . . . . . . . . . . . . . . 282 Cloning of PLCe . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 282 Splice Variants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 283 RA Domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 283 CDC25 Homology Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 284 Core Catalytic Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 284 Gbg Interaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 284 Regulation by GPCR Coupled with GaS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 285 Effects of Gene Knockout, Knockdown and a Lipase-Dead Mutant . . . . . . . . . . . . . . . . . . . . . . . . . . . . 285

5 5.1 5.2

Other Newly Isolated PLC Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 286 Sperm-Specific PLCz that Lacks the PH Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 286 Neuron-Specific PLCZ with C-terminal Extension . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287

6 6.1 6.2 6.2.1

PLC-Related Molecules Without the Catalytic Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 288 PRIP (PLC-L) as a PLCd1 Homologue Without Lipase Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 288 PRIP as a Regulator of Gabaergic Neuronal Signal Transduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 288 Future Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 289

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Abstract: Phosphoinositide-specific phospholipase C (PLC) comprises a family of multidomain phosphodiesterases. The family mediates signaling responses including the classically described conversion of PtdIns(4,5)P2 to Ins(1,4,5)P3, a Ca2+-mobilizing second messenger, and diacylglycderol, a protein kinase C-activating second messenger. It also mediates alterations in the intracellular localization and/or activity of a myriad of proteins that harbor phosphoinositide binding domains. Structural study of eukaryotic PLC has revealed that there is a prototypic structure for the enzyme represented by d-type isoforms. To date, at least 13 PLC isoforms and a few PLC-related molecules have been identified. Presumably, these molecules have evolved from the prototypic enzyme by addition or deletion of functional domains. By analyzing these modified structures, we can learn how efficiently the present isoforms exert diverse but specific physiological functions. This article gives an overview of the structures and roles of PLC isoforms and related molecules. List of Abbreviations: DAG, diacylglycerol; EGF, epidermal growth factor; EPAC, exchange protein activated by cAMP; ERK, extracellular signal-regulated kinase; EST, expressed sequence tag; GABARAP, GABAA receptor associated protein; GPCR, G protein-coupled receptors; LPA, lysophosphatidic acid; MAP, mitogen-activated protein; NES, nuclear export signal; NGF, nerve growth factor; RA, Respiratory acidosis; NLS, nuclear localizaiton signal; PDGF, platelet derived growth factor; PH, pleckstrin homology; PI3K, phosphatidylinositol 3-kinase; PIKE, PI3K enhancer; PKC, protein kinase C; PLC, phospholipase C; PP1a, protein phosphatase 1a; PRIP, PLC-related, but catalytically inactive protein; RGS, regulator of G protein signaling; RTK, receptor tyrosin kinase; SH, src homology; TIM, triose isomerase

1

Introduction

Phosphoinositide (PI) signaling pathways are ubiquitous regulatory systems in eukaryotic cells. They contribute to a myriad of processes, including signaling from cell surface receptors, regulation of the actin cytoskeleton, and intracellular trafficking. As a whole, they function as pivotal controllers in cell proliferation, differentiation, and even in cell death. In these pathways, it is thought that the breakdown of PtdIns(4,5)P2 by receptor-dependent activation of PI-specific phospholipase C (PLC: EC 3.1.4.11) at the plasma membrane plays an essential role, liberating lipid metabolites that transmit signals indirectly from the plasma membrane to targeted organelles, such as the endoplasmic reticulum and the nucleus. In addition to the role at the plasma membrane, growing evidence suggests that the intracellular translocation of PLC isoforms occurs during the cell cycle and in response to external stimuli. PtdIns(4,5)P2 is also a substrate for phosphatidylinositol 3-kinase (PI3K) and directly regulates a variety of cell functions including cytoskeletal reorganization, channel activity, exocytosis, and endocytosis. PtdIns(4,5)P2 homeostasis, therefore, is strictly regulated by PLC and other modifying enzymes.

2

PLC Isoforms

Since the late 1980s, multiple isozymes were purified and their genes were cloned. At present, the mammalian PLC superfamily has been identified as being composed of six major groups, PLCb, PLCg, PLCd, PLCe, PLCz, and PLCZ (> Figure 14-1). PLCa reported earlier has not been classified as a functional phospholipase because of the lack of homology with other isoforms and uncertainty in that it exerts specific phospholipase activity. At present, at least 13 isoforms with PtdIns(4,5)P2-hydrolyzing activity have been identified in the mammalian PLC family. In addition, two genes encoding a catalytically inactive phospholipase with a genetic alteration at the catalytic site have been reported and their neuronal function will be discussed later. The central paradigm is that PLC isoforms catalyze the hydrolysis of PtdIns(4,5)P2 to Ins(1,4,5)P3 and diacylglycerol (DAG) in response to the activation of cell surface receptors (Rhee and Bae, 1997; Rebecchi and Pentyala, 2000; Fukami, 2002). The isoforms contain X and Y domains, which form the catalytic core, and various regulatory domains such as the PH domain, which binds specifically to PtdIns(4,5)P2 and tethers the entire molecule to membranes. PLCs serve as regulatory molecules, reducing the amount of

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. Figure 14-1 Structures of mammalian PLC isoforms and related molecules. A schematic representation of domain organization for each isoform is shown. The core domain consists of the PH domain, EF hands, the catalytic domain (a TIM barrel structure incorporating the X and Y regions of highly conserved sequences) and the C2 domain. PLC-related catalytically inactive protein (PRIP or PLC-L) has a TIM barrel structure lacking the catalytic activity. The ligand for each PH domains is expressed. An approximate molecular size (as numbers of amino acid residues) and known binding protein(s) are listed besides the structure of each isoform family

PtdIns(4,5)P2 as well as producing the second messengers that activate both Ca2+ and protein kinase C (PKC) signaling. d-type isoforms are thought to be prototypic among PLC isoforms and are found in most eukaryotic cells, while the newly cloned PLCe and PLCZ, as well as the classical PLCb and PLCg subfamilies, are only found in metazoans (Song et al., 2001; Hwang et al., 2005), and PLCz has been detected only in sperm (Cox et al., 2002). > Figure 14-2 illustrates a dendrogram showing interrelationship among the PLC isoforms. The molecular size of the isoforms ranges from 70 kDa for the PLCz isoform to 230–260 kDa for PLCe. While PLCd is only 85 kDa, b- and g-type are in the range of 120–155 kDa. Lower eukaryotes such as yeast and slime mold contain only d-type isoforms, suggesting that d-type is the prototype and other types are evolved from archetypal PLCd.

3

Core Structure of PLC

With the sole exception of PLCz, the d-type isoforms show the simplest structures. The simplest eukaryote, yeast, has a single PLCd-like gene (Yoko-o et al., 1993; Andoh et al., 1995), suggesting that PLC-d is the prototypical isoform of PLC.

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. Figure 14-2 A dendrogram of PLC isoforms. The amino acid sequences of all human PLC isoforms and related molecules were aligned by ClustalW (http://clustalw.ddbj.nig.ac.jp/). Subfamilies of PLC and related molecules that share common functions are grouped by color. Note that the simplest PLC isoform is z-type, which shows limited expression pattern among the isoforms. PRIP (PLC-L) shows more similarity with d-type PLCs rather than h- or b-type, although they have C-terminal extensions in common. PLC« and PLCg1,2 may be diverged early from other isoforms

With regard to structures, all PLC isoforms contain, from N-terminus to C-terminus, the PH domain (with the exception of PLCz), the EF hand domains, catalytic X and Y domains connected by the X-Y linker domain, and the C2 domain. Although the general fold of these domains is well-conserved, their properties vary from one isoform family to another, or even among the same family. In addition to these conserved domains, each isoform possesses auxiliary domains that make it possible to exert specific intermolecular interactions. Examples of these domains are the src homology (SH) domains and a split PH domain in PLCg, the tail domain of PLCb and PLCZ, and the Ras-association (RA) domain and Ras-GTPase exchange factor (Ras-GEF)-like domain in PLCe. Since various isoforms have multiple additional domains, they could serve as both scaffold proteins functioning as platforms for signal transduction and as catalysts.

3.1 Structure of PLCd1 Since d-type PLC is a prototypical isoform of PLC, its structure has been explored in detail. X-ray crystallographic studies of two domains of PLCd1, the PH domain (Ferguson et al., 1995), and the rest of the isoform (Essen et al., 1996), in the mid 1990s provided a vital framework for understanding the conserved ‘‘core’’ structure of the PLC superfamily. Possible orientation of both the PH domain and the rest of the molecule of rat PLCd1 are shown in > Figure 14-3. PH domain: The PH domain of PLCd1 is composed of seven antiparallel b-sheets capped with a terminating N-terminal a-helix at one end. Binding of the ligand, either Ins(1,4,5)P3 or the head group of

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. Figure 14-3 The 3D structure of rat PLCd1. Possible orientation of both the PH domain and the rest of the molecule are shown. Since no structure of full-length PLCd1 is available, a composite of two crystal structures for the PH domain with Ins(1,4,5)P3 (PDB code 1MAY) and the rest of the molecule with Ins(1,4,5)P3 (PDB code 1DJX) were illustrated. The position of the PH domain therefore is arbitrary. Disordered regions such as the X-Y linker region and the N-terminus of EF hand domains are not described. In addition to the PH domain, which tether the whole enzyme to the PtdIns(4,5)P2 on the plasma membrane, there are at least two membrane targeting domain; one is a hydrophobic rim region near the catalytic pocket of the TIM barrel catalytic core, and the other is in the C2 domain. Ca2+ binding sites are found in the catalytic core and the C2 domain, and both are essential for the catalytic activity and the membrane targeting, respectively. In this structure, only one Ca2+ is shown in the C2 doamin, although the site can accommodate up to three Ca2+

PtdIns(4,5)P2, is orientated by the loops within the antiparallel b-sheet structure. The PH domain of PLCd1 determines the membrane localization of PLCd1 by anchoring the protein via a specific high affinity interaction with PtdIns(4,5)P2 (Garcia et al., 1995; Lemmon et al., 1995; Kavran et al., 1998; Varnai et al., 2002). This PtdIns(4,5)P2-dependent localization of the PH domain has been shown to provide an indirect regulatory mechanism of PLCd1 through regulation of the frequency of encounter between the PLCd1 core catalytic domain and its substrate, PtdIns(4,5)P2 (Lomasney et al., 1996; Yagisawa et al., 1998). Although most of the PLC isoforms contain the PH domain, the function of the PH domain in each isoform may differ. For example, the PH domain of PLCd1 utilizes the amphipathic a2-helix,

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which protrudes from the core b-sheet structure, to stabilize the membrane interaction by placing the hydrophobic surface of the helix against the hydrophobic portion of the membrane interface (Tuzi et al., 2003). The lipid microenvironment, such as the presence of an acidic lipid PtdSer or cholesterol, can alter the interaction (Uekama et al., 2007). This suggests that the PH domain of PLCd1 could serve as a sensor of the membrane environment to regulate the affinity of the PH domain for the membrane interface. Nevertheless, this kind of subtle fine-tuning function may not be conserved in the PH domains of other PLCs. The residues within the PH domain of PLCd1 that orientate the ligand-binding are not conserved in either PLCb or PLCe. In fact, the PH domain of PLCb isoforms do not possess the highly polarized electrostatic surface potential present in the PH domain of PLCd1 (Singh and Murray, 2003), and it has been reported that there is no apparent specific binding of the PH domain to phosphoinositides (Runnels et al., 1996; Tall et al., 1997). The binding specificity of the PH domain also varies. Many reports show that the N-terminus PH domain of g isoforms specifically binds PtdIns(3,4,5)P3 and serves to recruit g isoforms to membranes containing PtdIns(3,4,5)P3 (Falasca et al., 1998; Matsuda et al., 2001). Alternatively, the PH domain of the b isoform shows little affinity for phosphoinositides, although there is a report indicating that PLCb1 binds with Ptd(3)P through its PH domain and that this binding is necessary for the enzymatic function in cells in which PI3K is activated (Razzini et al., 2000). It is likely that, although the PH domain of the b-isoform PLC interacts with phospholipid bilayers, PtdIns(4,5)P2 is not the molecule that is responsible for its recruitment to the membranes. Catalytic core: The central catalytic core structure of PLC has been resolved by X-ray crystallographic studies on PLCd1 (Essen et al., 1997). The catalytic domain is formed from the X and Y domain, comprising 150 and 120 amino acid residues, respectively. The domain is composed of a TIM (triose isomerase) barrel structure with alternating a-helices and b-strands. This barrel structure is surrounded by two tandem clusters of EF hand motifs and a C-terminal C2 domain. The core structure of each of these modules is the most highly conserved among PLC isoforms. The X domain corresponds to the first lobe of the barrel, the Y domain forms the second one, and the residues surrounding the catalytic domain are on both lobes. The catalytic residues of the TIM barrel are located at one end of the barrel, and the catalytic site is formed in a small cavity composed of residues of both the X and Y domains. The site is surrounded by hydrophobic residues that penetrate into the phospholipid bilayers to facilitate membrane interaction and access to the substrate (Essen et al., 1997; Ellis et al., 1998). Since the PH domain is highly flexible and PtdIns(4,5)P2 is not always a good binding partner for PH domains of some isoforms, it is this hydrophobic rim on the surface of the TIM barrel structure that is directly responsible for introduction of the substrate into the catalytic core. All enzymatically active PLC isoforms require Ca2+ for catalytic function. Structural and mutational studies on PLCd1 have revealed that residues in the catalytic domain serve to orientate the substratebinding and also Ca2+ (Ellis et al., 1998). Four amino acid residues in PLCd1, each of which are also conserved in the PLCb isoforms, and PLCg isoforms are shown to be important for substrate recognition; Lys438 (X domain), Ser522 (Y domain), and Arg 549 (Y domain) are important for 4-phosphate recognition on the head group of phosphoinositides and Lys440 (X domain) for 5-phosphate recognition (Ellis et al., 1998). X-Y linker: The general function of the X-Y linker region of PLC isoforms is not known. The linkers of the PLCg subtype, because of the presence of two SH2 domains, a SH3 domain and a split PH domain, may be evolved to show distinct and additional functions as mentioned later. Other isoforms have rather short but flexible X-Y linker regions (whose disordered structure cannot be ‘‘seen’’ by the X-ray crystallographic studies of PLCd1), composed of clusters of basic and acidic residues. Since proteolytic cleavage of PLCd1 accompanied by cleavage between the X and Y regions increased basal PLC activity, it was suggested that the X-Y linker regulates the active site by direct occlusion (Ellis et al., 1993). A possible role for the X-Y linker domain to regulate substrate access to the catalytically active site also came from a study indicating that calmodulin stabilized conformation of the X-Y linker to occlude the active site and inhibit the activity of PLCd1 (Sidhu et al., 2005). Also, the basal specific activity of PLCg2 (Horstman et al., 1996) or PLCb2 (Zhang and Neer, 2001), whose X-Y linker domain was eliminated, increased, suggesting the regulatory function of this domain on control of the catalytic activity.

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Another possible function of the X-Y linker domain is in regulation of the nuclear import of some PLC isoforms. In the case of PLCd1, the NLS-like domain is located in a region spanning from the C-terminal of the X domain to the middle of the X-Y linker domain (Okada et al., 2002). Residues predicted to constitute a nuclear localization signal (NLS) are also present in the X-Y linker of PLCz (Larman et al., 2004). EF hand domains: Usually PLC isoforms contain two pairs of lobes of EF hand motifs, consisting of four helix-loop-helix structures. In the case of PLCd1, the domain does not bind Ca2+ but rather serves as a flexible link between the PH domain and the rest of the molecule, according to the X-ray crystallographic studies (Essen et al., 1997; Ellis et al., 1998). Therefore, after binding of the PH domain to its ligand, PtdIns (4,5)P2, the EF hand domains allow the catalytic core and the C2 domain to interact with the membrane PtdIns(4,5)P2. Direct binding of Ca2+ to the EF hand motifs is an issue of controversy. The conformation of the second lobe of the EF hand motif and the EF hand/C2 interface is similar to the Ca2+-saturated form of calmodulin bound to its target proteins (Essen et al., 1996). Therefore, it is unlikely that this region (the second lobe) works as a Ca2+ sensor. However, PLCd1 lacking the EF hand domain lost Ca2+ sensitivity dramatically (Nakashima et al., 1995). Moreover, a biochemical study revealed that Ca2+ binds to the EF hand domain of PLCd1 and the bound Ca2+ is necessary for the efficient interaction of the PH domain with PtdIns(4,5)P2 (Yamamoto et al., 1999). In the case of PLCd1, a canonical leucine-rich nuclear export signal (NES) sequence, which is functional, was found localized in residues 164–177 of the first lobe of the N-terminal EF hand domain (Yamaga et al., 1999). There is accumulating evidence indicating that some PLC isoforms are imported into the nucleus or shuttle between the cytoplasm and the nucleus. Although there have been several NLS sequences or NLS-like sequences found in PLC isoforms; sequences homologous to canonical NES are only found in the EF hand motifs (Yagisawa et al., 2006) (> Figure 14-4). C2 domain: The C2 domains of PLC isoforms are composed of 120 residues. Structural studies of PLCd1 revealed the presence of three Ca2+ binding sites in this domain (Essen et al., 1996; Essen et al., 1997; Grobler and Hurley, 1998). The C2 domain of PLCd1 consists of eight antiparallel b-strands and three loops at one end of the b-sandwich structure to form the Ca2+ binding site: loop1 (642–653), loop2 (675–680), and loop3 (706–714). Deletion of the C2 domain of PLCd1 decreased its activity, suggesting that Ca2+ acts as an element recruiting the catalytic core domain adjacent to the C2 domain to a membrane that contains acidic phospholipid such as PtdSer. Actually, PLCd1 forms a functional complex with PtdSer and Ca2+ in vitro (Lomasney et al., 1999). The aforementioned Ca2+-binding region in the C2 domain is conserved in d-type PLC but not in the b- and g-type isoforms. It is not known whether the Ca2+-binding characteristic is major function among PLC isoforms, since the PLCb1 C2 domain can bind the GTP-bound form of the Gaq molecule (Wang et al., 1999) in concert with the C-terminal extension domain. > Figure 14-5 illustrates a ‘‘tether and fix’’ model (Katan and Williams, 1997) in which these domains regulate membrane association and dissociation of PLC, taking PLCd1 as an example. The catalytic core consists of EF, TIM barrel and C2 domains is recruited to the membrane by the function of the PH domain and fixed to the plasma membrane. A hydrophobic rim near the catalytic pocket (see > Figure 14-3) is inserted to the hydrophobic portion of the plasma membrane. In the presence of Ca2+, the C2 domain also stabilizes the interaction between the catalytic core and the inner leaflet of the plasma membrane. This ‘‘fixed’’ state assures the access to the substrate PtdIns(4,5)P2, causing an efficient hydrolysis of it. A rapid decrease in the amount of PtdIns(4,5)P2 at the membrane surface results in a shift in a binding equilibrium between the membrane and PLCd1. PLCd1 rapidly dissociates from the membrane to become inactive.

3.2 PLCd Subfamily 3.2.1 Yeast PLC Regulates Cell Growth Both budding and fission yeasts (Saccharomyces cerevisiae and S. pombe) possess only one PLC of the d type, Plc1p, encoded by a gene, PLC1 (Yoko-o et al., 1993; Andoh et al., 1995). Budding yeasts that lack the gene or have mutations in the catalytic domains of Plc1p display multiple problems when stressed, although there is little effect in unstressed conditions (Flick and Thorner, 1993; Yoko-o et al., 1993). They show

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. Figure 14-4 Regions of PLCd1 important for the nucleocytoplasmic shuttling. Schematic representation of the rat PLCd1 structure is shown with the nuclear export signal (NES) sequence of PLCd1. The corresponding regions in other PLCd isoforms and a consensus leucine-rich NES sequence, where ‘‘X’’ denotes any amino acid and bold letters indicate important hydrophobic residues, are also shown. Polybasic regions corresponding to the C-terminal of the X domain and X-Y linker region is necessary for the nuclear localization of PLCd1. Corresponding regions of various PLCs are also displayed for comparison. Sequences of PLC isoforms without species names are those of rat. Adapted from (Yagisawa et al., 2006)

growth defects in hyperosmotic conditions (Flick and Thorner, 1993), in synthetic media (Yoko-o et al., 1993; Flick and Thorner, 1998), at high temperatures (Yoko-o et al., 1995), display defective cytokinesis (Payne and Fitzgerald, 1993), unusual sensitivity to UV-irradiation (Andoh et al., 1998), and vacuole fragmentation (Seeley et al., 2002). The Plc1p of budding yeast is found in the nucleus and is localized to centromeric loci at the G2/M checkpoint, affecting kinetochore function, possibly by modulating the structure of centromeric chromatin (DeLillo et al., 2003).

3.2.2 Mammalian PLCd Subfamily The mammalian PLCd subfamily was originally reported as being composed of four isozymes, PLCd1–4. However, since bovine PLCd2 was found to be a homologue of human/mouse PLCd4, the PLCd family is now considered to be composed of three isoforms, d1, d3, and d4 (Irino et al., 2004).

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. Figure 14-5 A regulation model of PLCd1. The PH domain binds to PtdIns(4,5)P2 at the plasma membrane. In the presence of Ca2+ the catalytic core move towards the inner leaflet of the plasma membrane, since (1) the C2 domain forms firm interaction with acidic phospholipids including PtdIns(4,5)P2, and ii) a hydrophobic rim region in the vicinity of the catalytic site is inserted into the membrane. When substrate PtdIns(4,5)P2 is hydrolyzed, the resultant second messenger Ins(1,4,5)P3 is released into the cytoplasm and binds to IP3R of ER membranes. The decrease in the levels of PtdIns(4,5)P2, and increase of Ins(1,4,5)P3 in the vicinity of inner leaflet of the plasma membrane dissociate the enzyme from the plasma membrane and move it into the cytoplasm. This ‘‘tether and fix’’ model, originally claimed by Katan and Williams (1997), elucidates the PH domain- and Ca2+dependent PtdIns(4,5)P2 hydrolysis of PLCd1. PLCd1 can be activated by heterodimeric G protein (Gh)-linked G protein-coupled receptors (GPCR) or directly by a RhoGAP called p122RhoGAP (DLC1). Phosphorylation levels of PLCd1 may also modify the catalytic activity. In addition to the role at the plasma membrane and in the cytoplasm, PLCd1 plays role in the nucleus, since it binds to importin b, a carrier protein for nuclear import, in a Ca2+-dependent manner and shuttles between the nucleus and cytoplasm (Yagisawa, 2006)

PLCd1 is expressed abundantly in most tissues (Lee et al., 1999). In normal rat heart, PLCd1 is more abundant than the PLCb1 and PLCg1 isoforms (Hwang et al., 2004). Because of the presence of the PH domain that shows a high affinity for PtdIns(4,5)P2 (Kd of 10–100 nM), PLCd1 is generally distributed at the inner leaflet of the plasma membrane and in the cytoplasm of various cell types. Although there are only a few reported agonists whose receptors directly activate PLCd1, a myriad of external stimuli that modulate the levels of PtdIns(4,5)P2 can dissociate PLCd1 from the plasma membrane. Loss of PtdIns(4,5)P2 by activation of any of the PLC isoforms or of PtdIns(4,5)P2 phosphatases can facilitate the dissociation of PLCd1. The dissociation can be readily observed after elevation of intracellular Ca2+ levels, since PLCd1 shows high sensitivity to Ca2+. It is unlikely, however, that phosphorylation of PtdIns(4,5)P2 via activation of PI3K to produce PtdIns(3,4,5)P3 reduces the membrane-attached PLCd1. In some cell types, translocation of PLCd1 to perinuclear regions was observed after dissociation from the plasma membrane (Fujii et al., 1999). PLCd3 is expressed at high levels in the rodent heart, aorta, and esophagus and weakly in the testis, retina, and adipose tissue (Enomoto et al., 2005). The catalytic and regulatory properties of PLCd3 are similar to those of PLCd1 (Pawelczyk and Matecki, 1998). PLCd3 also requires Ca2+ for its activity, but shows a relatively low activity in cellular conditions that fully activate PLCd1 (Pawelczyk and Matecki, 1998). The activity of PLCd3 is stimulated by polyamines and by basic proteins such as protamine and histone in vitro. PLCd3 binds to lipid membranes via the PH domain, and phosphatidic acid supports this binding through Ca2+-dependent interaction with the C2 domain (Pawelczyk and Matecki, 1999). In rat

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liver, PLCd3 is present in both the membrane and cytosolic fraction and is absent in the nuclear fraction (Pawelczyk and Matecki, 1998). PLCd4 has been cloned as a cell-growth-related gene from a cDNA library prepared from regenerating rat liver (Liu et al., 1996) and also from normal rat brain (Lee and Rhee, 1996). The substrate specificity and Ca2+ dependency of PLCd4 were similar to those of PLCd1 (Lee and Rhee, 1996). In highly proliferating cells such as regenerating liver cells, hepatomas, and src-transformed cells, the mRNA levels of PLCd4 were much higher than those in resting cells (Liu et al., 1996). It has been reported that there are three alternatively spliced variants of PLCd4, termed Alt I, II, and III, whose expression levels vary from tissue to tissue (Lee and Rhee, 1996; Nagano et al., 1999).

3.2.3 Gene Knockout of d Isoforms Nakamura et al. have produced PLCd1-deficient mice (Nakamura et al., 2003). The mice show epidermal hyperplasia of skin cells and undergo progressive hair loss. Their keratinocytes and skin cells show impaired intracellular Ca2+ elevation and PKC activation. In addition, they develop spontaneous skin tumors (Nakamura et al., 2003). The knockout mice have a slight phenotypic difference from the wild type, showing a defect in the normal development of skin but overall cell division and proliferation are similar to that in wild-type mice (Nakamura et al., 2003). The skin of PLCd1 knockout mice displays the typical inflammatory phenotype, including increased dermal cellularity, leukocyte infiltration, and expression of proinflammatory cytokines (Ichinohe et al., 2007). Moreover, overexpression of PLCd1 attenuates LPSinduced production of the proinflammatory cytokine IL-1, suggesting that PLCd1 regulates homeostasis of the immune system in the skin (Ichinohe et al., 2007). PLCd3 gene-deficient mice did not show particular phenotypic differences from wild-type mice (Enomoto et al., 2005). Important roles of PLCd4 in the testis, where the isoform is predominantly expressed, have been clarified by production of PLCd4 gene knockout mice (Fukami et al., 2003). PLCd4 is required for intracellular Ca2+ mobilization and sustained Ca2+ increase through store-operated channels in the acrosome reaction in sperm (Fukami et al., 2003). Moreover, the initiation of DNA synthesis, as well as the appearance of PKCa/bII in the S phase is delayed in regenerating livers from the PLCd4 knockout mice, suggesting that PLCd4 controls the timing of DNA synthesis in liver generation (Akutagawa et al., 2005). None of the gene knockout mice of d-type PLC was embryonic lethal, suggesting the functional redundancy among the isoforms during the development of mice. In fact, Nakamura et al. reported that PLCd1 and PLCd3 double knockout (DKO) embroys died during mid-gestation, with a severe disruption in placental development, suggesting the close functional interrelationship between the two isoforms in embryogenesis (Nakamura et al., 2005). To dissect the physiological roles of the isoforms, it would be useful to carry out gene knockin experiments of various mutants to embryonic fibroblasts (MEF) from knockout mice for each d isoform to see whether any alterations in phenotypes occur.

4

Isoform Families that have Auxiliary Domains: Regulation Under the Control of Specific Molecular Interaction

4.1 PLCg with the SH2/SH3-Containing X-Y Linker Domain PLCg isozymes are characterized by the presence of a linker domain containing two SH2 domains plus one SH3 domain (SH223 domain) sandwiched by a split PH domain between the X and Ydomains. Polypeptide growth factors, such as platelet derived growth factor (PDGF), epidermal growth factor (EGF), and nerve growth factor (NGF) activate PLCg isoforms to produce Ins(1,4,5)P3 and DAG from PtdIns(4,5)P2 in various cell types via activation of their respective receptors, which are intrinsic protein tyrosin kinases. Binding of the growth factor to its receptor results in dimerization of receptor subunits and stimulates intrinsic protein tyrosine kinase activity, introducing autophosphorylation on the specific tyrosine residues in the intracellular moiety of the receptor molecule. The autophosphorylation sites create high-affinity docking sites for the SH2 domains of various effector proteins including PLCg and PI3K. For example,

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PLCg1 recognizes and binds to Y1021 of the PDGF receptor (b-chain) (Claesson-Welsh, 1994), Y766 of the FGF receptor, and Y785 of the NGF receptor (Obermeier et al., 1993). The role of the two SH2 domains were dissected using mutants of each domain of PLCg1 in fibroblasts. The results showed that it is the N-terminal SH2 domain that is responsible for binding to the receptor and for transduction of signaling (Ji et al., 1999; Poulin et al., 2000). Phosphorylation of bovine PLCg1 by growth factor receptors (PDGF, EGF, FGF, or NGF) occurs at the identical site: tyrosine residues 771, 783, and 1254 (Kim et al., 1990). Substitution of Y783 to F (phenylalanine) resulted in a lack of response of PLCg1 to PDGF but association of the enzyme with the PDGF receptor was not changed (Kim et al., 1991). Interestingly, mutation of some of the aforementioned autophosphorylation sites on the growth factor receptors, which serve as the PLCg1 recognition sites, prevents the receptor association of PLCg1 and Ca2+ signaling that follows receptor activation, but tyrosine phosphorylation of PLCg1 still occurs. Therefore, two signals, i.e., tyrosine phosphorylation of the PLCg1 and association of the enzyme with the receptor via the N-terminal SH2 domain of the X-Y linker are necessary for proper growth factor-induced activation of PLCg1. To recruit activated PLCg in the vicinity of the membrane area where the substrate PtdIns(4,5)P2 is located, another machinery for recruitment of PLCg1 seems to be present. Activation of the receptor protein kinase also generates PtdIns(3,4,5)P3 via activation of PI3K. An inhibitor of PI3K drastically downregulates both Ins(1,4,5)P3 production and Ca2+ mobilization (Bae et al., 1998). PtdIns(3,4,5)P3 binds both the N-terminal PH domain (Falasca et al., 1998) and C-terminal SH2 domain of PLCg1 (Falasca et al., 1998; Rameh et al., 1998). Therefore, translocation of PLCg1 to the membrane, where it hydrolyzes the substrate PtdIns(4,5)P2, is also facilitated by growth factor-mediated production of PtdIns(3,4,5)P3, in concert with recruitment of the molecule toward the growth factor receptor upon activation. PLCg1 can be a facilitator of guanine nucleotide exchange (GEF) activity via the SH3 domain in the SH223 region. In fact, the SH3 domain of PLCg1 associates with SOS1 through its proline-rich region and stimulates its GEF activity (Kim et al., 2000). Furthermore, results from Ye et al. (Ye et al., 2002) describe the ability of PLCg to facilitate GEF for the recently identified GTPase, PIKE (for PI3K enhancer) (Ye et al., 2000), localized in the nucleus. The SH3 domain, rather than the catalytic domain, of PLCg is required for aiding PIKE. Interestingly, the mitogenic activity of PLCg depends neither on its phospholipase activity nor on its N-terminal SH2 domain-dependent receptor association, but rather on its interaction with PIKE. Thus, although PLCg participates in cellular mitogenesis, this evidence indicates that the catalytic activity of PLCg is not essential to the process (Wang and Moran, 2002). The SH3 domain of PLCg1 also has a special role in controlling neurite outgrowth and neuronal differentiation in relation to NGF signaling. It was shown recently that overexpression of PLCg1 inhibits neurite outgrowth and prolonged proliferation. PLCg1 alters cell cycle regulatory proteins such as PCNA and cyclin D1, such that the cells exit from cell growth arrest. Deletion of the SH3 domain or the entire SH223 domains, but not deletion of the N-terminal SH2 domain alone or of both the N-SH2 and C-SH2 domains, abolishes the differentiation-inhibitory effects of PLCg1. The proliferation activity of PLCg1 via its SH3 domain may be related to growth arrest by NGF, inhibiting neuronal differentiation (Nguyen et al., 2007). In the X-Y linker domain of PLCg1, the SH223 structure is sandwiched by a split PH domain. Unlike the N-terminal PH domain that shows specific affinity for PtdIns(3,4,5)P3 (and b-tubulin, see below), the split PH domain of PLCg1 binds PI(4)P and PI(4,5)P2 (Chang et al., 2002). The structure of the N-terminal half of the split PH domain is responsible for orientation of binding of these ligands (Kim et al., 2004). Moreover, a binding partner of the split PH domain of PLCg1 was identified as translational elongation factor (EF)-1a, which is a GTP-binding factor and a phosphatidylinositol 4-kinase activator (Chang et al., 2002). The binding of EF-1a seems to control the binding of PtdIns(4,5)P2 to the split domain, although PtdIns(4,5)P2 does not bind to EF-1a directly. PLCg1 is activated in the presence of EF-1a, but a PLCg1 mutant lacking EF-1a binding is not, suggesting that the elongation factor regulates PLCg1 signaling via the split PH domain. Recently, both the N-terminal and the split PH domain of PLCg were shown to bind to b-tubulin using a pull-down assay of NIH3T3 cells (Chang et al., 2005). PLC-g1 and b-tubulin colocalized in the perinuclear region in COS-7 cells and cotranslocated to the plasma membrane upon agonist stimulation, suggesting that PLCg is recruited to the membrane in a form associated with microtubules. Indeed, PLCg1 is highly concentrated in mitotic spindle fibers and involved in its formation. It seems likely that PLCg1 modulates microtubule assembly by b-tubulin via its N-terminal PH domain and the split PH domain, and vice versa; b-tubulin facilitates PLCg1 activity (Chang et al., 2005).

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In summary, SH2, SH3, and a split PH inserted in the X-Y spanning domain are key components in the mechanism of activation (> Figure 14-6); PLCg1 binds to receptor tyrosine kinases (RTKs) through the N-terminal SH2 domain, leading to tyrosine phosphorylation of PLCg1 and activation of the enzyme. The SH3 domain is involved in GEF activity for nuclear PIKE. The two PH domains have distinct functions; the N-terminal domain interacts with PtdIns(3,4,5)P3 and mediates plasma membrane localization, whereas the split PH shows affinity for PtdIns(4,5)P2, PI(4)P and binding proteins that control PLC activity and intracellular localization. Details of the regulation of PLCg by RTKs and other typrosine kinases have been intensely studied and reviewed (Rhee and Bae, 1997; Rebecchi and Pentyala, 2000; Rhee, 2001) (> Figure 14-7). . Figure 14-6 Mitogenic signaling by PLCg. Generally, growth factor stimulation triggers autophosphorylation of receptor tyrosine kinase (RTK), which functions as docking sites for many SH2 domain-containing proteins. One of them, phosphatidylinositol 3-kinase (PI3K) converts PtdIns(4,5)P2 to PtdIns(3,4,5)P3, which are the target for both the N-terminal PH domain and the C-SH2 domain. PLCg is phosphorylated on a certain tyrosine residue and undergoes substantial conformational changes. This conformational changes stabilizes the interaction, letting the enzyme stay in the proximity to the plasma membrane, stimulating conversion of PtdIns(4,5)P2 to Ins(1,4,5)P3 and diacylglycerol (DAG). DAG then stimulates the downstream PKC/MAPK signaling cascade. PLCg1 binds b-tubulin and associates with microtubule. This result suggests that intracellular trafficking of PLCg1 by microtubule-related cytoskeletons. Another hallmark of recent findings is that the SH2 domains and the PLC activity are not necessary for NGF-induced mitogenesis (Ye et al., 2002). PLCg1 translocates from cytoplasm to nucleus in response to the growth factor and it act as a GEF for PIKE, a GTPase and an activator for nuclear PI3K, via its SH3 domain. The classical PKC/MAP signaling cascade and the PIKE-GEF signaling via the SH3 domain need not be mutually elusive. It has not been elucidated yet how PLCg1 enters the nucleus in response to growth factors or whether association to the receptor is necessary or not. The SH3 domain of PLCg1 also binds Sos1 through proline-rich domain

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. Figure 14-7 Regulation of PLC by heterotrimeric G proteins and small G proteins. (a) PLCb isoforms are regulated by direct interaction with both Gbg and GTP-bound form of Ga (Ga*) subunits of the Gq family of heterotrimeric G proteins. Activated (GTP-bound) forms of other small G proteins are also described with asterisks. (b) PLC« isoform possess a GEF domain that activates Ras and Rap isozymes. These activated GTPases directly stimulate the PLC activity of PLC« through direct binding to the RA domains. Ras is also activated downstream of RTK. Rap is activated by GPCR by activation of exchange protein activated by cAMP (EPAC) (see text). Ga12/13 can activate Rho by Rho-GEF such as p115-Rho-GEF and activated Rho binds to the catalytic core of PLC« and stimulate enzymatic activity in a ‘‘Y-insert’’-dependent manner. Modified from (Harden and Sondek, 2006)

4.2 PLCb Under the Control of Heterotrimeric G Proteins PLCb isoforms are thought to transduce signals from heterotrimeric G proteins. The standard G protein model of PLCb activation is that binding of the ligand triggers receptor catalyzed exchange of GTP for bound GDP on the a subunit. The GTP-charged subunit then dissociates in the plane of the membrane, and either the a subunit monomer or the bg-heterodimer, or both, bind to PLCb, increasing its catalytic activity, thereby amplifying the initial receptor stimulus.

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4.2.1 Gbg Dimer Activates PLC Isoforms PLCb isoforms function as effector enzymes for the 7-transmembrane receptor superfamily, namely G protein-coupled receptors (GPCR) (Ji et al., 1998). The bg heterodimers of heterotrimeric G proteins that couple with the receptors are recognized as regulators of many effectors including PLCb. The Gbg dimer strongly activates PLCb2 and PLCb3, whilst PLCb1 is weakly activated when subunits are reconstituted into artificial and biological membranes (Camps et al., 1992), and PLCb4 is insensitive (Lee et al., 1994). Binding of PLCb2 occurs through multiple sites on the PLC structure. The site of interaction of PLCb2 with Gbg was originally mapped to a narrow region in the Y domain (Sankaran et al., 1998), but later to the PH domain (Wang et al., 2000). Thus, Gbg can have different effects on the function of PLCb; one possibility is that the tethering role of the PH domain to the membrane is modulated. The region of the Gbg subunit that interacts with effector molecules such as PLCb and adenylate cyclase was shown to overlap with that for binding of Ga subunits (Panchenko et al., 1998). This may explain why heterotrimeric G proteins, after activation, efficiently act as activators of the effector such as PLCb; bg dimers and the GTP-bound form of a subunits activate PLCb independently.

4.2.2 PLCb with C-terminal Extension (tail) Domain When compared with the structure of d-type PLC, b-type PLC is slightly more complex, since it contains the tail (or C-terminal extension) domain, consisting of 400 amino acid residues that cannot be found in the d-type or g-type isoforms. The domain plays important roles in membrane binding through acidic lipids, nuclear localization (putative NLS sequences are present in the domain), and activation by the Ga subunit of heterotrimeric G proteins (Kim et al., 1996; Jenco et al., 1997). Deletion of this domain inhibited Gaq-mediated activation of PLCb1, but not the catalytic activity itself (Park et al., 1993). PLCb also utilizes the core structure of PLCd, but has evolved in such a way that the PH domain of PLCb2 binds to the GTP-bound form of small G protein Rac and Gbg or the C-terminal extension of all b-type isoforms to bind to Gaq (> Figure 14-7a).

4.2.3 Expression and Distribution of PLCb Isoforms PLCb1 is widely expressed in the brain; this isoform is prominent in the pyramidal cells of the hippocampus and cerebral cortex (Ross et al., 1989; Kim et al., 1997). PLCb1 exists as alternative splice variants b1a and b1b, both of which are abundant in the brain, but with b1a more predominant than b1b (Bahk et al., 1994). Like PLCg2, the expression of PLCb2 is restricted to hematopoietic cells and functions in leukocyte signaling (Lee et al., 1996; Li et al., 2000). PLCb3 is expressed in a wide variety of tissues but is mainly distributed in the brain (Jhon et al., 1993). The highest levels of expression of b3 mRNA were found in cerebellar Purkinje and granule cells and in the pituitary gland (Tanaka and Kondo, 1994). PLCb4 was first isolated from brain cerebellum (Min et al., 1993) and the retina (Lee et al., 1993) and high levels of its mRNA were detected in cerebellar Purkinje and granule cells (Jiang et al., 1996; Kim et al., 1997). It should be noted that the fly PLCb4 homologue, NorpA, was identified as an eye-specific gene and the results strongly suggests that PLCb4 is involved in the visual signaling process (> Figure 14-7b).

4.2.4 Gene Knockout Studies of PLCb Isoforms The role of PLCb1 in the brain was also investigated by generating gene knockout mice (Kim et al., 1997). It was found that PLCb1/ mice developed epilepsy causing sudden death after birth, suggesting that PLCb1 is involved in signal transduction in the cerebral cortex and hippocampus. Since carbachol-induced PtdIns (4,5)P2 hydrolysis was markedly inhibited, the PLCb1 signal seems to couple predominantly to the muscarinic acetylcholine receptors. The authors also investigated the brain function of PLCb4/ mice

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and found that they showed ataxia, suggesting that PLCb4 works through the metabotropic glutamate receptor in the cerebellum. Mice lacking PLCb3, when compared with the wild type, exhibited up to a tenfold decrease in morphine sensitivity as detected by production of antinociception (Xie et al., 1999). m-Opioid regulation of voltagesensitive Ca2+ channels in primary sensory neurons (dorsal root ganglion) was apparently attenuated in PLCb3-null mice. PLCb3 constitutes a significant pathway involved in negative modulation of m-opioid responses. Since in neurons, activation of PKC with phorbol esters markedly reduces the m-opioid-induced Ca2+ current, this negative modulation is probably due to PLC activation. A study of mice lacking PLCb2 and PLCb3 revealed that the signaling pathways have an important role in chemoattractant-mediated production of superoxides and regulation of protein kinases, but not chemotaxis (Li et al., 2000). Mice lacking PLCb4 were impaired in their visual processing ability (Jiang et al., 1996).

4.3 PLC«: Under the Control of Small G Proteins and Heterotrimeric G Proteins The recent discovery of a new type of PLC, PLCe, with multiple auxiliary domains has altered the concept of G protein-stimulated cell signaling (Wing et al., 2003a; Bunney and Katan, 2006; Harden and Sondek, 2006). It was previously thought that G protein-mediated PLC activation was segregated into those mediated by heterotrimeric G proteins versus those promoted by the small GTPases of the Ras superfamily. PLCe exhibits a unique regulatory potential, both as initiator and recipient of activated G protein signals.

4.3.1 Cloning of PLC« In 1996, a database study detected an open reading frame containing two Ras-associating (RA) domains, the TIM barrel fold of PLC isoforms and a putative N-terminal Ras-GEF-like (CDC25) domain (Ponting and Benjamin, 1996). Then Kataoka and colleagues isolated a new PLC, PLC210, from Caenorhabditis elegans as a binding protein for LET-60, the worm Ras ortholog (Shibatohge et al., 1998). This 210 kDa protein possesses the conserved catalytic core domain (PH, EF, X/Y-TIM barrel, and C2 domains) of other PLC isoforms. The human ortholog of PLC210 was cloned thereafter and shown to contain several unique domains such as the C-terminal RA domain and the N-terminal CDC25 domain, indicating that PLC activity can be a downstream target of small G proteins and also a modulator of small G proteins in addition to its function as a PtdIns(4,5)P2-hydrolyzing enzyme (Kelley et al., 2001; Lopez et al., 2001; Song et al., 2001).

4.3.2 Splice Variants There are several splice variants of human PLCe. Two splice variants (PLCe1a and PLCe1b) are derived from the human PLCe1 gene. PLCe1a and PLCe1b have a similar potential to be stimulated by diverse signaling pathways via tyrosine kinase and GPCRs and both can function as an effector of Ras. The expression pattern shows broader mRNA expression of PLCe1a in normal tissues; furthermore, in most cell lines expressing PLCe, PLCe1a is the only splice variant present. A demethylating agent recovers expression of PLCe1a and PLCe1b suggesting epigenetic silencing of PLCe through hypermethylation. Expression levels of PLCe were greatly reduced in tumor tissues. The inhibitory effects of PLCe on cell viability and proliferation were seen in a cell line that has a silenced PLCe expression (Sorli et al., 2005).

4.3.3 RA Domains Sequence analysis revealed that there are two tandem RA domains (RA1 and RA2) in the C-terminus. RA domains in general consist of 100 amino acid residues with low sequence homology, and interact directly

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with Ras family GTPases (Ponting and Benjamin, 1996). 1H, 13C, and 15N NMR studies resolved the characteristics of the two RA domains (RA1 and RA2) from PLCe (Harris et al., 2005). The presence of tandem RA domains is a unique feature among Ras effectors. The two domains are structural homologues and the presence of tandem domains may be due to DNA duplication early in the course of evolution (Bunney et al., 2006). Kataoka’s group first demonstrated GTP-dependent binding of H-Ras to the RA domain of PLC210 in vitro (Shibatohge et al., 1998), and then showed that Ras and Rap1 were associated with the RA2 domain of human PLCe. Although both RA1 and RA2 interact with H-Ras and are indispensable for the activation of PLCe, RA2 shows a higher affinity for H-Ras (Kelley et al., 2001). The RA2–Ras interaction seems strictly regulated by their conformation, since point mutations in H-Ras and single amino acid substitutions (e.g., K2150E) in the RA2 domain of PLCe abrogated the GTP-dependent interaction between Ras and the RA2 domain (Kelley et al., 2001). TC21, Rap1A, Rap2A, and Rap2B stimulate PLCe in an RA2-dependent manner similar to H-Ras (Kelley et al., 2004). Coexpression of an activated H-Ras mutant with PLCe induces its translocation from the cytosol to the plasma membrane (Song et al., 2001). Upon stimulation with EGF, similar translocation of ectopically expressed PLCe is observed, which is inhibited by coexpression with dominant-negative H-Ras. When PLCe is coexpressed with Rap1, it translocates to a perinuclear compartment such as the Golgi apparatus. Activation of the EGF receptor may recruit GEFs to membranes to form the GTP-bound form of H-Ras or Rap1, resulting in the recruitment of PLCe to the plasma membrane or to the Golgi membrane, respectively. The structure of PLCe RA domains (RA1 and RA2) were analyzed by NMR and that of the RA2/Ras complex by X-ray crystallography (Bunney et al., 2006). Despite the similarity between the ubiquitin-like folds of RA1 and RA2, only RA2 binds to Ras. Some of the features of the RA2/Ras interface are unique to PLCe, while the ability to make contact with both the switch I and II regions of Ras is shared only with PI3K. It seems that the RA2 domain, in a mode specific to PLCe, has a role in membrane targeting with further regulatory effect on PLC activity.

4.3.4 CDC25 Homology Domain The CDC25 homology domain in general has a GEF function for Ras family GTPases. PLCe contains a CDC25 homology domain and an adjacent REM (Ras exchange motif) domain in the N-terminus (Kelley et al., 2001; Lopez et al., 2001; Song et al., 2001). Therefore, it is intriguing to examine if PLCe works as a Ras-GEF to activate Ras. Jin et al. used an in vitro assay to study the specificity of PLCe-Ras-GEF activity. PLCe has an activity toward Rap1 but not to Ras, Rap2A, or Rho. It was concluded, therefore, that a pivotal role of the CDC25 homology domain was in amplifying Rap1-dependent signal transduction, including the activation of PLCe itself, at specific subcellular locations such as the Golgi apparatus (Jin et al., 2001). These results differ from those of Lopez et al. demonstrating that a lipase-dead mutant of PLCe retains the capacity to function as a GEF for Ras. They also showed that mitogen-activated protein (MAP) kinase (or extracellular signal-regulated kinase, ERK), a downstream effector of Ras, can be activated by PLCe or by its CDC25 homology domain alone. In their hands, Ras activation attenuates PLCe activity. The exact specificity (Rap1 or H-Ras) of the CDC25 homology domain is still unclear, but the downstream effector kinases such as B-Raf, MEK, and MAPK are activated via this domain. Thus the relationship between Ras/Rap1 activation and PLCe activation seems complex. Using cells expressing a PDGF receptor mutant incapable of activating PLCg, which makes the phosphoinositide/Ca2+ response more complex, Song et al. observed persistent PDGF-mediated activation of ectopic PLCe in hematopoietic BaF3 cells (Song et al., 2002). They proposed that a rapid initial phase of activation is mediated by Ras and a prolonged phase is promoted by Rap1. PDGF-induced proliferation of cells lacking the PLCg response is dependent on the activation levels of PLCe, suggesting a critical role of PLCe in cell survival and proliferation.

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4.3.5 Core Catalytic Domain Constitutively active Ga12 and Ga13, but not Gaq, enhances the enzymatic activity of PLCe (Lopez et al., 2001; Wing et al., 2001). This is rather surprising, since most PLCb isoforms respond to a subunit of the Gq family. In PLCe lacking CDC25, the PH and RA domains can still be activated by Ga12 and Ga13. There is no direct evidence that Ga12 or Ga13 binds PLCe. Rather, it is possible that Rho is activated by Ga12/13 via the action of Rho-GEF, and activated Rho has been postulated to interact with PLCe (Wing et al., 2003b). A candidate targeting region of Rho binding is in the catalytic core, i.e., a unique 60–70-residue insert (residues 1,667–1,728 of rat PLCe, in some reports denoted as ‘‘the Y-insert’’) in the Y domain. A mutant of PLCe lacking this region is no longer activated by Ga12/13, or Rho (Wing et al., 2003b; Seifert et al., 2004), suggesting that the region is responsible for transducing signals from Ga12/13-coupled GPCRs. Ga12/13, as well as Rho, Rac, and Ral can therefore stimulate PLCe in an RA2-independent manner (Kelley et al., 2004). Regarding Ga12/13- and Rho-dependent activation of PLCe, Hains et al. explored the downstream signals of lysophosphatidic acid (LPA) and thrombin receptors (Hains et al., 2006). Receptor-mediated activation of PLCe was inhibited by coexpression with the RGS (regulator of G protein signaling) domain of p115-Rho-EF, a GTPase-activating protein for Ga12/13 but not by expression of the RGS domain of GRK2 (G protein-coupled receptor kinase 2), which inhibits Gaq signaling. Conversely, activation of neither the Gq-coupled M1 muscarinic nor the P2Y2 purinergic receptor was enhanced by coexpression with PLCe. These studies illustrate that specific LPA and thrombin receptors promote inositol lipid signaling via activation of Ga12/13 and Rho.

4.3.6 Gbg Interaction In addition to Ga, the Gbg dimer interacts with PLCe. Expression of Gb1g2 in Cos-7 cells stimulates the enzymatic activity of PLCe (Wing et al., 2001). The combination of bg subunits is interchangeable, so that Gb1g3 or Gb2g2 can promote the stimulation. The stimulation of PLCe by Gb1g2 is not altered when a mutation in the RA2 domain that eliminates the effect of Ras is introduced, suggesting that Ras–RA2 interaction is independent of Gbg activation.

4.3.7 Regulation by GPCR Coupled with GaS Schmidt et al. (2001) describe a mechanism through which GPCR coupled with Gas stimulates PLCe. Activation of the adenylate cyclase-coupled b2-adrenergic receptor expressed in HEK-293 cells or the endogenous receptor for prostaglandin E1 in N1E-115 neuroblastoma cells induced Ca2+ mobilization and PLCe stimulation caused by cAMP formation. These responses are mediated by a Rap GTPase, specifically Rap2B, activated by a GEF named EPAC (exchange protein activated by cAMP) regulated by cAMP. Rap2B directly binds to the RA2 domain of PLCe in a GTP dependent manner (Song et al., 2002). Gs-coupled receptor stimulation of PLCe via formation of cAMP and activation of Rap2B was also confirmed by Evellin et al. (2002). Thus, Ins(1,4,5)P3 generation and the Ca2+ response of GPCRs to catecholamines and prostaglandins are stimulated by EPAC.

4.3.8 Effects of Gene Knockout, Knockdown and a Lipase-Dead Mutant Studies to explore overall function of PLCe have been conducted in gene knockout and knockdown experiments as well as by introduction of an enzyme lacking PLC activity. So far various phenotypes linked to fragmented function have appeared. Ovulation in C. elegans is dependent on the Ins(1,4,5)P3 signaling pathway activated by the RTK LET-23. Deletion mutants of the gene plc-1, a nematode ortholog of PLCe, show a novel ovulation

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phenotype whereby oocytes are trapped in the spermatheca due to delayed dilation of the spermathecauterine valve (Kariya et al., 2004). A PLCe deficient mouse strain was created to explore the role of PLCe in cardiac function and pathology (Wang et al., 2005). Isolated cardiac myocytes from PLCe/ mice have decreased b-adrenergic receptor-regulated Ca2+ transient amplitudes and the mice are more susceptible to hypertrophy in response to chronic cardiac stress, suggesting regulation by a pathway involving camp, EPAC, and Rap. Thus PLCe may be involved in heart development and function. An important role of PLCe in cardiac development was also confirmed using a mouse model system in which lipase activity was catalytically inactivated by gene targeting (Tadano et al., 2005). In the lipase-dead mice, malformation of aortic and pulmonary valves, a defect in valve remodeling at the late stages of semilunar valvulogenesis, was observed. The role of PLCe in chemical carcinogen-induced skin tumor development has also been investigated using PLCe/ mice. The mice are less susceptible to the onset of skin tumors than their wild-type counterpart; development of malignancy was retarded in the skin of PLCe/ mice, although the activation stage of H-ras was similar in both PLCe/ backgrounds (Bai et al., 2004). Therefore, PLCe plays a crucial role in Ras oncogene-induced carcinogenesis. Phorbol ester-induced hyperplasia was also impaired in PLCe/ mice suggesting a link with downstream effectors of PKC. A new physiological function for PLCe in the regulation of integrin activation has been reported (Lad et al., 2006). In some effector mutants H-Ras can suppress integrin activation in an ERK-independent manner. These mutants synergize with wild-type PLCe to suppress integrin, reducing cell adhesion, but a lipase-dead PLCe mutant prevented this suppression. Moreover, knockdown of endogenous PLCe with siRNA resulted in blockage of the mutant H-Ras-mediated integrin suppression. These results show that H-Ras suppresses integrin affinity via both Raf/MEK/ERK-dependent and PLCe-dependent signaling pathways. A recent study showed that PLCe is a novel effector of R-Ras, an atypical member of the Ras subfamily of small GTPases that enhances integrin-mediated adhesion and signaling. R-Ras coprecipitated with PLCe and increased its activity. Knockdown of PLCe with short interfering RNA reduced the formation of ruffling lamellipodia in R-Ras-expressing cells. R-Ras signaling regulates the organization of the actin cytoskeleton to sustain membrane protrusion through the activity of PLCe (Ada-Nguema et al., 2006). Using RNA interference to knock down PLC isoforms in Rat-1 cells, PLCe was shown to predominantly account for sustained rather than early PtdIns(4,5)P2 hydrolysis downstream of GPCR receptors for endothelin (ET)-1, LPA, or thrombin. Conversely, it was suggested that PLCb3 is responsible for acute responses to agonists for these receptors (Kelley et al., 2006). It is interesting to note that a naturally occurring mutation in the human genome causes an early-onset nephrotic syndrome. A missense mutation in an exon encoding the PLCe catalytic domain results in the histological characteristics of focal segmental glomerulosclerosis (Hinkes et al., 2006). In summary, as expected from the diverging signaling cascades both upstream and downstream of PLCe, deletion of PLCe or destruction of its enzymatic activity results in a variety of phenotypes. Therefore, many reports so far give only a fragmental picture of the functions of PLCe, which are either linked to, or independent of, Ras or other small G proteins.

5

Other Newly Isolated PLC Isoforms

5.1 Sperm-Specific PLCz that Lacks the PH Domain PLCz may represent the physiological stimulus for egg activation and development in mammalian fertilization (Swann et al., 2004; Swann et al., 2006). The sperm activates development by causing a prolonged series of intracellular Ca2+ oscillations that are generated by increased production of Ins(1,4,5)P3. No PLC isoforms, however, have been detected as a candidate for a sperm factor that triggers the oscillations. PLCz has recently been cloned from a mouse testis expressed sequence tag (EST) database as the cytosolic sperm factor that triggers Ca2+ oscillations in eggs at fertilization (Saunders et al., 2002). Expression or injection of recombinant PLCz into eggs triggers Ca2+ oscillations and induced development up to the blastocyst stage. This PLC isoform, with a molecular size of 70 kDa, is the smallest of all the mammalian PLC isoforms

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and shows a distinctive structure lacking the PH domain. PLCz, and the closely related PLCd1, have similar Km values for PtdIns(4,5)P2, but PLCz is around 100 times more sensitive to Ca2+ than PLCd1 (Kouchi et al., 2004; Nomikos et al., 2005). This means that the Ca2+ levels of resting cells (100 nM) are sufficient for its activity. The third EF hand domain of PLCz is responsible for this high Ca2+ sensitivity (Kouchi et al., 2005). Northern blot analysis revealed that the expression of the isoform only occurs in spermatids. PLCz is a major PLC in the sperm of the mouse, hamster, pig, and human (Cox et al., 2002; Saunders et al., 2002) and is responsible for Ca2+ oscillation during fertilization utilizing mitochondrial ATP in the oocytes (Dumollard et al., 2004; Rogers et al., 2004). This isoform has an NLS sequence in its X-Y linker domain. Dissection of the cell cycle (M-phase) dependent Ca2+ oscillations and disruption studies of the NLS sequence in mouse embryos suggests that nuclear sequestration of PLCz is one of the major controlling mechanisms for the cell cycle-dependent regulation of Ca2+ oscillations following fertilization (Larman et al., 2004). Using a series of domain-deletion constructs, it was suggested that substrate specificity is retained in the structure of the catalytic TIM barrel, but the EF hand domains or C2 domains are indispensable for the very fine Ca2+ sensitivity of the enzyme (Nomikos et al., 2005). These results suggest that PLCz is evolved as specific machinery for the fine-tuning of Ca2+ in the first step of the cell cycle at very beginning of fertilization in oocytes. The absence of the PH domain in its structure may be a reflection that it is unnecessary for membrane targeting before (presumably in the sperm cytosol) and during the fusion step between the sperm and egg plasma membranes, since PLCz must readily diffuse into the egg cytoplasm. The precise localization of PLCz in the sperm, however, has not been established. It is paradoxical that expression of PLCd1 that lacks the PH domain did not cause Ca2+ oscillations in the egg (Saunders et al., 2002). Whether this is simply because of the difference in sensitivity to Ca2+ or because the remaining parts of PLCz can target the appropriate source of PtdIns(4,5)P2 in the eggs is unknown. A recent study by Nomikos et al. suggested that a basic cluster (374–385) in the X-Y linker domain of PLCz could not only anchor the protein to the membrane but also enhance the local concentration of PtdIns(4,5)P2 adjacent to the catalytic domain (Nomikos et al., 2007). Therefore, mechanisms for utilizing PtdIns(4,5) P2 by PLCz may be quite different from those for PLCd1 and other PLC isoforms.

5.2 Neuron-Specific PLCh with C-terminal Extension Data base information suggested the existence of a new type of PLC. In 2005, a PLC gene family designated as PLCZ emerged (Cockcroft, 2006). Searching the nucleotide sequence and EST database of mouse and human resulted in two closely related DNA sequences mapping to the human genome at positions 1p36.32 and 3q25.31 (Hwang et al., 2005; Nakahara et al., 2005; Zhou et al., 2005). The first identified gene (mapped to human genome 3q25.31) encodes 1,002 (human) and 1,003 (mouse) residues, and was denoted as PLCZ(1) (Hwang et al., 2005). Genomic analysis revealed that there could be two splice variants with 1,655 and 1,693 amino acids in humans. PLCZ1 has an apparent molecular size of 115 kDa, comprising the PH domain, EF hand domains, the TIM barrel core domain, the C2 domain, and, additionally, a long C-terminal sequence like b-type PLCs. The recombinant protein exhibited Ca2+-dependent PtdIns(4,5)P2 hydrolyzing activity. Northern blot analysis and protein blot analysis demonstrated that 115 kDa PLCZ1 is expressed mainly in nervous tissues especially in the brain and spinal cord. In situ hybridization revealed that PLCZ1 is abundantly expressed in the cerebral cortex, hippocampus, zona incerta, and cerebellar Purkinje cell layer (Hwang et al., 2005). The second identified gene was PLCZ2. Nakahara et al. reported that murine PLCZ2 consists of 1164 residues with a molecular size of 125 kDa (Nakahara et al., 2005). Like PLCZ1, PLCZ2 consists of a conserved canonical PLC core structure, but has an additional 290 amino acids at the C-terminus, resembling the situation with PLCb. However, sequence comparison showed a similarity to d-type isoforms. The PH domain of PLCZ2 functions as a membrane anchoring domain like that of PLCd1. Mouse PLCZ2 was much more sensitive to Ca2+ than PLCd1, whose maximal PLC activity was obtained in the presence of 10–100 mM Ca2+. The maximal PLC activity of PLCZ2 was obtained at 1 mM Ca2+ and it was sensitive to Ca2+ concentrations as low as 10 nM. Like PLCZ1, protein blot and Northern blot analyses

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proved that expression of PLCZ2 was restricted to nervous tissues, particularly the brain, predominantly in the hippocampus, cerebral cortex, and olfactory bulb. Zhou et al. reported that there are splice variants within the C-terminus both in the case of PLCZ1 and PLCZ2 based on in silico database cloning of PLCZ (Zhou et al., 2005). For example mouse PLCZ2 has five predicted splice variants and all transcripts were found in tissues including eye, brain, and lung. Some C-terminal splice variants of human and mouse PLCZ1 and PLCZ2 have expanded C-terminal domains ending with a class II PDZ domain binding motif, LLRL (Zhou et al., 2005). It is thought that this motif serves to interact with PDZ domain-containing proteins such as the Na+/H+ exchanger regulatory factor, similar to PLCb containing PDZ domain binding motifs. An immunoblot study of human astrocytoma cells indicates that a splice variant with a molecular size of 155 kDa is the natively expressed variant (Zhou et al., 2005). Coexpression studies using one of the PLCZ2 splice variants that shows maximum PLC activity in COS-7 cells demonstrated that, unlike coexpression with PLCb2 or PLCe, the PLC activity is not affected in the presence of Rac1, Rac2, or Rac3 (that activate PLCb2 activity), or RhoA, RhoB, or RhoC (that activate PLCe activity). No activation was observed following coexpression with the Ga subunits. Nontheless, coexpression of the Gbg dimer (Gb1g2) with PLCZ2 in COS-7 cells resulted in an increase in cellular PLC activity, suggesting the existence of an interaction between Gb1g2 and PLCZ2. The results obtained for the structure and function of the PLCZ family so far still need to be organized because of the presence of several splicing variants for each isoform. Probably, PLCZ1 is expressed specifically in the brain, whereas variants of PLCZ2 show a broader distribution (kidney, retina, brain, lung, etc.) apart from the fact that some are expressed mainly in the central nervous system. In addition to the higher sensitivity to Ca2+, at least a part of these isoenzymes can be activated by the Gbg subunit of heterotrimeric G proteins. Since PLCZ could generally be neuronal, potential roles that could not be substituted by other PLC isoforms in the brain have been proposed. It may be involved in the regulation of ion channels and ion transporters, since PtdIns(4,5)P2 is known to regulate the activation and desensitization of transient receptor potential (TRP) channels (Rohacs et al., 2005), a specific function that d-type isoforms cannot perform.

6

PLC-Related Molecules Without the Catalytic Activity

6.1 PRIP (PLC-L) as a PLCd1 Homologue Without Lipase Activity Investigation of chemically synthesized Ins(1,4,5)P3 analogs has led to the isolation of a binding protein with a molecular size of 130 kDa, characterized as a molecule with similar domain organization to PLCd1 but lacking the enzymatic activity (Kanematsu et al., 1992; Kanematsu et al., 1996) (for review (Kanematsu et al., 2005)). An isoform of the molecule was subsequently identified, and these molecules have been named PRIP (PLC-related, but catalytically inactive protein). Although amino acid identity between the major domains of PRIP and PLCd1 are approximately between 40 and 50%, the molecule is larger than the PLCd isoforms and unique regions are present at both the N-terminus preceding the PH domain and at the C-terminus. The reason why this molecule does not show PLC activity came from the amino acid alignments of the core TIM barrel domain: the residues within the catalytic domain critical for PLC activity (Glu341 and His356) in PLCd1 are not conserved in PRIP. Kohno et al. cloned a human gene homologue to rat PRIP and named it PLC-L (Kohno et al., 1995). Otsuki et al. later isolated a cDNA from mouse brain that encodes a protein with 64% sequence identity to PLC-L, and designated the original PLC-L as PLC-L1 and the new one as PLC-L2 (Otsuki et al., 1999). These two classes of genes are now called PRIP1 and PRIP2, respectively. Human PRIP2 (KIAA1092) was also found in the database and even C. elegans has a similar gene (Koyanagi et al., 1998), suggesting that this gene family diverged early from other PLC isoforms and holds distinct roles as PLC-like proteins without the catalytic activity. All PRIP family proteins have amino acid substitutions on the critical residues in the catalytic domain so that the proteins cannot hydrolyze PtdIns(4,5)P2. Since PRIP has an N-terminal PH domain that has strong affinity for Ins(1,4,5)P3 (Takeuchi et al., 1996), it could be involved in the regulation of inositol phosphate signaling rather than signaling of inositol

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lipid in the membrane. In fact, neither PRIP1 nor PRIP2 show strong membrane localization. Rather, PRIP1 sequesters Ins(1,4,5)P3 that mobilizes Ca2+ from the endoplasmic reticulum, since Ins(1,4,5)P3dependent Ca2+ release in permeabilized COS-1 cells is inhibited by the addition of the isolated PH domain of PRIP1 (Takeuchi et al., 2000). Moreover, agonist (bradykinin or EGF)-induced Ca2+ increase in cells overexpressing PRIP1 was dramatically reduced (Takeuchi et al., 2000). Sequestration of Ins(1,4,5)P3 by PRIP1 affects not only Ca2+ mobilization from the ER but also metabolism of inositol phosphate; Ins(1,4,5) P3-5-phosphatase activity and -3-kinase activity were also inhibited by the PRIP1 PH domain in a dosedependent manner (Takeuchi et al., 2000). Establishment of PRIP1 knockout mice (Kanematsu et al., 2002) made the aforementioned simple Ins(1,4,5)P3-sequester model a little more complex. Cultured neurons prepared from PRIP1 knockout mice responded less potently to the external stimulus (ATP) (Harada et al., 2005). Either up- or downregulation of PRIP1 may modify the fate of the Ins(1,4,5)P3 produced. In the latter case, trace amounts of PRIP1 in the vicinity of the plasma membrane could protect Ins(1,4,5)P3 from being hydrolyzed by Ins(1,4,5)P3-5phosphatase localized at the inner surface of the membrane. These contradicting results suggest the importance of the physiological levels of PRIP1 in the regulation of Ins(1,4,5)P3 and Ins(1,4,5)P3-mediated Ca2+ responses.

6.2 PRIP as a Regulator of Gabaergic Neuronal Signal Transduction A recent search for binding partners of the PRIP family revealed its close relationship with the regulation of a neurotransmitter receptor. Yeast two hybrid screening of a brain cDNA library using the N-terminal region of PRIP1 as the bait identified GABARAP (GABAA receptor associated protein) and PP1a (protein phosphatase 1a) as binding partners (Yoshimura et al., 2001). Both PRIP1 and PRIP2 have a consensus sequence for the binding of PP1a and the binding and activation of PP1 depends on phosphorylation of the threonine residue in the PP1a binding site (Terunuma et al., 2004). GABAA receptors are ligand-gated chloride channels and important for fast synaptic inhibition in the central nervous system. GABARAP binds GABAA receptors and is thought to be responsible for clustering of GABAA receptors at the postsynaptic membranes. PRIP binds to GABARAP and inhibits the association of GABARAP with the g2 subunit of GABAA receptors (Kanematsu et al., 2002). PRIP1 is expressed predominantly in the central nervous system, while PRIP2 shows broad tissue distribution, including the brain. It is suggested that GABAA receptor modulation by associating molecules is dependent on the phosphorylation stage. Results indicating that PRIP binds directly to PP2A (Kanematsu et al., 2006), PP1a, and the intracellular loop of the b-subunits of GABAA receptors (Terunuma et al., 2004) support the idea that PRIP plays an exclusive function in regulation of the GABAA receptor. Kanematsu et al. generated PRIP1 and PRIP2 double knockout (DKO) mice and analyzed GABAA receptor function in relation to the action of benzodiazepines from the electrophysiological and behavioral aspects. They demonstrated that the sensitivity to the drug was reduced in DKO mice suggesting the alteration of g2 subunits of GABAA receptors in these mice (Kanematsu et al., 2002; Mizokami et al., 2007). Pharmacological analyses showed the number of the surface diazepam-binding sites in DKO mice was reduced. The association between GABAA receptors and GABARAP was reduced significantly in DKO mice neurons. It is implicated that PRIP works as a scaffold protein for GABARAP: by binding with the b-subunit of GABAA receptor it delivers GABARAP to the g2 subunit of the receptor, eventually causing the translocation of the receptor complex in secretory vesicles to the cell membrane. Thus, apart from its role as a sequestering molecule for Ins(1,4,5)P3, PRIP works as an important regulatory molecule in neuronal signal transduction, regulating the trafficking of multisubunit nurotransmitter receptors (Kanematsu et al., 2007).

6.2.1 Future Perspective Although the 3D structure of the d isoform has been analyzed, the structure of the b, g, and e isoforms have not been elucidated in depth. So X-ray crystallographic analyses of these isoforms will provide important

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information concerning the mode of intramolecular interaction and regulation of domain-mediated signal transduction. The dynamics of membrane–PLC interaction have not been resolved, although, under limited conditions, we can now visualize the kinetics of the protein–protein interaction at a single molecule level. Emerging evidence suggests that PLC isoforms are general targets of not only heterotrimeric G proteins but also small G proteins. PLCe is a typical example of this, but many ‘‘classical’’ PLC isoforms are also targets of small G proteins. For example, PLCb2 activity and its mode of interaction with membranes is regulated by Rac2 (Illenberger et al., 2003). Moreover, when CaM binds to a novel IQ-type CaM-binding motif within the catalytic region, inhibition of PLCd1 activity that could be reversed by Ral can be observed (Sidhu et al., 2005). So, even PLCd1 is under the regulation of small G proteins. Thus, studies of PLC regulation have now entered into a new era where small G proteins act as machinery for most of the isoforms. To obtain the overall physiological function of each isoform, further gene knockout and knockdown experiments should be conducted and carefully dissected.

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Phospholipid Signaling and Cell Function

Y. Nozawa

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298

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General Properties of Phospholipase D . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298

3 3.1 3.2 3.3 3.4

Regulation of Phospholipase D . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 299 Protein Kinase C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 300 Small GTPase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 300 Protein Tyrosine Kinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 301 Mitogen-Activated Protein (MAP) Kinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 302

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Functions of Phospholipase D . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 303

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Role of Phospholipase D Signaling in Neural Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 305 Neurotransmitter Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 305 Differentiation (Neurite Outgrowth) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 305 Survival (Antiapoptosis) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 306 Neurodegeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 307

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Abstract: Phospholipase D (PLD) is an important member of the phospholipid signaling enzymes, which catalyzes generation of phosphatidic acid (PA), a second messenger, and choline, a precursor of the neurotransmitter acetylcholine (ACh). PA can then be converted to 1,2-diacylglycerol (DAG) by PA phosphohydrolase and to lysoPA (LPA) by phospholipase A2 which act as protein kinase C activator and as an extracellular agonist, respectively. In addition to the role of PA as a signaling mediator, it plays a biophysical role as a membrane perturbant for fusion pore formation in exocytosis. The PLD activation is linked to a wide array of physiological and pathophysiological cell responses, including the rapid responses such as secretion, vesicle trafficking, cytoskeletal rearrangement, and the long-term responses such as proliferation, differentiation, survival, apoptosis, and degenerative disorders. List of Abbreviations: ACh, acetylcholine; ActD, ActinomycinD; AD, Alzheimer’s disease; aPKC, atypical PKCs; bAPP, bA precursor protein; cPKC, conventional PKC; DAG, 1,2-diacylglycerol; dbcAMP, dibutyryl cyclic AMP; DGK, DAG kinase; Egr-1, early growth response-1; ERK, extracellular signal-regulated kinase; 4E-BP1, 4E binding protein1; LPA, lysoPA; LPC, lysophosphatidylcholine; MAP, Mitogen-Activated Protein; MEK, MAP kinase-ERK kinase; mTOR, mammalian target of rapamycin; nPKC, novel PKC; p38MAPK, p38 MAP kinase; PAP, phosphatidic acid phosphohydrolase; PA, phosphatidic acid; PC, phosphatidylcholine; PH, pleckstrin homology; PI(4,5)P2, phosphatidylinositol 4,5-bisphosphate; PIs, phosphoinositides; PKC, protein kinase C; PLC, phospholipase C; PLD, Phospholipase D; PTK, protein tyrosine kinase; PTP, tyrosine phosphatase; ROCK, Rho kinase; SCAMP2, secretory carrier membrane protein2; TGN, trans-Golgi network; tPA, tissue plasminogen activator

1

Introduction

Phospholipase D (PLD) was first discovered as a phospholipid-specific phosphodiesterase by Hanahan and Chaikoff (1947) in carrot extracts and has been found to be widespread in the plant kingdom. However, it is now well known that PLD is widely distributed in a variety of species, including animals, plants, fungi, protozoa, and bacteria. This enzyme in mammalian tissues hydrolyzes phosphatidylcholine (PC) to phosphatidic acid (PA) or choline. PA can then be converted to 1,2-diacylglycerol (DAG) by phosphatidic acid phosphohydrolase (PAP) and lysoPA (LPA) by phospholipase A2. DAG produced by PAP can be reconverted to PA via phosphorylation by DAG kinase (DGK). These PLD-mediated metabolites act as lipid second messengers. The interplay of these enzymes controls the levels of PA, DAG, and LPA. PA has been thought to play a role in signal transduction and cellular functions. DAG is a crucial factor for activating protein kinase C (PKC), and LPA acts as an agonist through specific receptors. Abundant evidence has shown that PLD is activated by various stimuli, such as growth factors, neurotransmitters, hormones, chemoattractants, and oxidants. PLD is thus recognized to be an important phospholipid-signaling enzyme (Exton, 1947; Cockcroft, 2001; McDermott et al., 2004; Jenkins and Frohman, 2005; Cazzolli et al., 2006). PLD-mediated signal transduction has also been observed in a variety of brain- and neural-derived cells, including primary neurons and glial cells, as well as neuroblastoma, glioblastoma, astrocytoma, and pheochromocytoma cell lines. Substantial experimental evidence has indicated that PLD signaling is implicated in physiology and pathophysiology of the neural system (Klein et al., 1995; Klein, 2005).

2

General Properties of Phospholipase D

PLD has a unique catalytic reaction, transphosphatidylation. When a primary alcohol such as butanol or ethanol is present as a nucleophilic donor, PLD catalyzes transesterification (transphosphatidylation), by which phosphatidylalcohol is produced at the expense of PA. Isobutanol is reactive but secondary and tertiary butanols are not. The production of phosphatidylalcohol, which is not rapidly metabolized, is often used as a potential parameter for the PLD activity.

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The mammalian PLD family consists of two isoforms, PLD1 (120 kDa, 1074 amino acids) and PLD2 (105 kDa, 933 amino acids) (Liscovitch et al., 2000; Jenkins and Frohman, 2005). These isoforms are 5% identical and differ in their N and C termini. PLD1 has a 116-amino acid loop region domain. An alternatively splicing variant of PLD1b that lacks 38 amino acids within the loop region between the catalytic domains is very similar to PLD1a in its catalytic properties, e.g., requirement of phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) and responsiveness to Arf and Rho family proteins, and PKC (Hammond et al., 1997). PI(4,5)P2 is also an essential cofactor of PLD2 activity. PLD belongs to the superfamily that includes, in addition to the hydrolytic enzyme PLD, cardiolipin/ phosphatidylserine synthase, Yersinia toxin, and poxvirus envelope protein. Most members of the PLD superfamily are dimeric structures containing the highly conserved sequence HKD motif (HxxxxKxD: H, histidine; x, any amino acid; K, lysine; D, aspartic acid) (McDermott et al., 2004). PLD1 and PLD2 have two HKD motifs that are essential for the enzyme activity. The HKD motifs undergo intramolecular dimerization to form a catalytic site. In addition to the intramolecular dimerization, an association between two molecules of PLD has been shown by the coimmunoprecipitation experiments (Kam and Exton, 2002); the differentially tagged PLD1 proteins coimmunoprecipitate, indicating the dimerized complex formation. It is also shown that PLD1 coimmunoprecipitates with PLD2, suggesting the formation of a heterodimer between different PLD isoforms. Other highly conserved regions of PLD1 and PLD2 are the pleckstrin homology (PH) domain and the phox homology (PX) domain in the N-terminal region. The PX domain has been proposed to be involved in protein–protein interaction and in binding various phosphoinositides (PIs), such as PI(4,5)P2, PI(3,4)P2, PI(3,4,5)P3. Recently, PI(3,4,5)P3 was found to specifically bind to the PH domain of PLD1 (but not PLD2) and stimulate its activity (Lee et al., 2005). The PH domain is thought to participate in translocation and regulation of PLD activity via interaction with PI(4,5)P2. The PH domain shows a relatively weak but selective affinity for PI(4,5)P2. Mutation of this domain in PLD2 causes its redistribution from the plasma membrane to early endosomes (Sciorra et al., 2002). It is also observed that the PH domain-mutated PLD2 is catalytically active but unresponsive to stimuli. This finding implies a critical role of the PH domain in differential translocation of PLD that is important for PLD function. Thus, PIs have a dual role in PLD regulation; membrane targeting and activation. PI(4,5)P2 also binds to a polybasic PIP2-binding motif with the catalytic core. Another unique structure is the loop region (116 amino acids) that is present in PLD1 but not PLD2. Although the exact function remains unknown, it may act as a negative regulatory component (Sung et al., 1999). As for subcellular localization, it has been generally acknowledged that PLD1 localizes to the internal membranes, such as Golgi apparatus, endoplasmic reticulum, secretory vesicles, and endosomes, whereas PLD2 appears to be associated with the plasma membrane. Such preferential distribution of PLD isoforms can be explained at least in part by the PIs binding with their PH and PX domains. Both PLD1 and PLD2 are reported to be palmitoylated and phosphorylated on serine or threonine residues under basal conditions. These modifications are not required for the activity but may affect membrane association (Exton, 2002).

3

Regulation of Phospholipase D

The relative distribution of PLD1 and PLD2 is distinct depending on the cell type. The activities of these two PLD enzymes exhibit different requirement for their catalytic activity. PLD1 has a low basal activity and is regulated by many other factors besides the PIs described above, namely by PKC, Arf and Rho family proteins, and tyrosine kinase (Exton, 1998; Cockcroft, 2001; Exton, 2002; McDermott et al., 2004). In contrast, PLD2 has a high basal activity, but can be stimulated moderately by Arf and PKC. However, the mechanism for the high basal activity of PLD2 remains unknown. Both PLD1 and PLD2 show a stringent requirement for PI(4,5)P2, indicating a close association with PIP5-kinase which generates PI(4,5)P2 from PI(4)P. This represents a positive feedback loop involving PLD and PIP5-kinase.

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3.1 Protein Kinase C Many isozymes have been identified in mammalian cells, and are classified into three subgroups consisting of the classical conventional PKC (cPKC) (a, bI, bII and g; Ca2+ and PMA dependent), the novel PKC (nPKC) (d, e, y;, Z; Ca2+ independent and PMA dependent), and the atypical PKCs (aPKC) (z, l); Ca2+ and PMA independent). It has been known for a long time that exposure of various cells to agonists or PMA stimulates PLD activity, indicating involvement of PKC (Exton, 2002; Nozawa, 2002; Becker and Hannun, 2005). Furthermore, prevention of agonist-induced PLD activation by PKC inhibitors or downregulation of PKC by prolonged treatment with PMA, and overexpression or knockdown of PKC isozymes provide further evidence that PKC plays a role in PLD activation. However, the regulatory mechanism(s) for PLD activation by PKC has not been fully clarified. Several lines of evidence have indicated that PKCa activates PLD1 through a phosphorylation-independent mechanism, with a protein–protein interaction being proposed as the major mechanism for PKCa-mediated PLD activation. The isolated membrane-bound PLD is activated by PMA in the presence of PKCa and PKCb without ATP-dependent phosphorylation in HL60 cells (Ohguchi et al., 1996). Hu and Exton (2003) have demonstrated that both the regulatory and catalytic domains are required for PLD activation and that a residue in the C terminus of PKCa (Phe663) is essential for binding, activation, and phosphorylation of PLD1. Another study shows that the PLD1 is dually regulated through both phosphorylation and protein interaction with PKCa, and probably by PKCb in vivo (Oka et al., 2002). On the other hand, phosphorylation has been suggested to be involved in the downregulation of PLD1 activity at the later stage after agonist or PMA treatment. This is also true with PLD2; interaction, rather than phosphorylation, is responsible for PLD2 activation by PKCa, and phosphorylation contributes to the inactivation of the enzyme (Chen and Exton, 2004). In contrast, some studies have shown evidence to denote a more pivotal role of phosphorylation in PLD regulation. PKCa has been reported to phosphorylate PLD1 on Ser 2 and Thr 147 in the PX domain, and Ser 561 in the negative regulatory loop region, and mutation of these residues reduces PMA-induced PLD1 activity (Kim et al., 1999). It has also been shown that direct phosphorylation of PLD2 by PKCd is important for the regulation of PLD2 activity in PC12 cells (Han et al., 2002).

3.2 Small GTPase There is a large body of evidence indicating importance of several small GTPases, such as Arf and its subtypes, Rho family proteins, and RalA in stimulation of PLD1 activity (Exton, 1998; McDermott et al., 2004). PLD1a and PLD1b are highly responsive to activation by Arf1 and can also be activated by purified Rho family proteins, RhoA, Rac1, and Cdc42 (Hammond et al., 1997). In contrast, PLD2 shows no significant response to the latter proteins but is modestly responsive to Arf1, suggesting that PLD2 is activated by Arf under specific conditions. There are six subtypes of the Arf family, which activate PLD1 with modest differences in potency and efficacy. Myristoylated forms of these proteins are more potent for activation of PLD1, because of their efficient membrane association. Activation of PLD1 by Arf proteins is caused by their direct interaction. The site on PLD1 that interacts with Arf has not yet been identified, though regions in the middle and the N terminus of PLD1 have been suggested. A possible indirect mechanism that has been proposed for PLD activation by Arf is PIP5-kinase dependent. Stimulation of cells with agonists induces translocation of cytosolic Arf to membranes where it can recruit several proteins including PIP5-kinase which produces PI(4,5)P2 from PI(4)P. Taken together, Arf1and Arf6 act as the active regulators for both PLD and PIP5-kinase, whereas PA produced by PLD activates PIP5-kinase and PI(4,5)P2 produced by PIP5-kinase activates PLD (Oude Weernink et al., 2007). Thus, the concerted cross talk between Arf, PLD, and PIP5-kinase, together with their products (PA, PI(4,5)P2), regulates the levels of these second messengers in the cell. Substantial evidence has been accumulating that RhoA, Rac1 and Cdc42 activate PLD1, but not PLD2 (Exton, 2002; McDermott et al., 2004). RhoA is involved in stress fiber formation, Rac1 in membrane ruffling, and Cdc42 in filopodia formation. Activation of PLD by direct interaction with these Rho family

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proteins is shown in vitro using purified PLD1 enzyme. The interaction site of RhoA has been demonstrated to be a basic region in the C terminus within residues 946–962 of PLD1. Several other sites have also been observed in the C terminus. The PLD1 interaction site on RhoA is thought to be within the switch 1 activation loop. In addition to such direct activation of PLD1 by Rho, an indirect Rho-dependent mechanism may be involved in PLD1 stimulation. By inactivation by monoglucosylation of RhoA GTPase with bacterial toxins (C. difficile toxin B, C. botulinum C3), the cellular PI(4,5)P2 level is decreased and PLD stimulation is prevented, thereby suggesting a PIP5-kinase-mediated PLD activation, as observed in the case of Arf. It has been proposed that PIP5-kinase and PLD activities are regulated through the action of Rho kinase (ROCK) (Oude Weernink et al., 2007), but the precise mechanism remains to be established. Another Rho effector, the PKC-related protein kinase N (PKNa, PKNb) has been demonstrated to directly interact with PLD1; PKNa binds to residues 228–598 and PKNb to residues 1–228 and 228–598 of PLD1 (Oishi et al., 2001). PKNa stimulates PLD1 activity in the presence of PI(4,5)P2 in vitro, whereas PKNb showed a modest stimulation. Ras does not directly stimulate PLD activity, but it mediates activation of PLD induced by v-src in vivo. RalA, a member of the Ras subfamily, is also thought to be a regulator of PLD activation in several cell types. This protein functions downstream of Ras and can directly interact with PLD. RalA may function in cooperation with Arf and Rho proteins.

3.3 Protein Tyrosine Kinase There are two types of protein tyrosine kinase (PTK); receptor type and nonreceptor (cytosolic) type. Growth factors such as EGF and PDGF and other agonists such as IgE and carbachol have been shown to activate PLD (Kumada et al., 1993; Schmidt et al., 1994; Ito et al., 1997a; Slaaby et al., 1998; Choi et al., 2004), but no clear evidence is available for direct phosphorylation of the enzyme by the tyrosine kinase activity of their receptors. Such receptor-mediated PLD stimulation is thought to occur through a signaling cascade involving PKC and Ras/Ral (Voss et al., 1999). On the other hand, there has been considerable evidence to suggest the involvement of nonreceptor PTK such as the Src family in PLD activation (Nozawa, 2002). Most evidences derive from the observation that PLD activation is inhibited by PTK inhibitors such as genistein and herbimycin. Furthermore, the fact that inhibition of tyrosine phosphatase (PTP) by vanadate can activate PLD or enhance its activity in combination with oxidant provides additional evidence for implication of PTK. Bourgoin and Grinstein (1992) first showed that stimulation with peroxyvanadate induces accumulation of tyrosine-phosphorylated proteins and concomitant activation of PLD in HL60 cells. It has also been demonstrated that in response to stimulation by peroxyvanadate PLD1 is tyrosine phosphorylated (Marcil et al., 1997). However, this apparent correlation between tyrosine phosphorylation and activation of PLD1 could not provide strong evidence for direct activation of PLD1 by tyrosine phosphorylation. Initial study with H2O2 demonstrated that it causes activation of PLD, which is independent of PKC and calcium in bovine pulmonary endothelial cells (Natarajan et al., 1996). The effects of the PTK inhibitors and PTP inhibitor vanadate on H2O2-induced PLD activation and protein tyrosine phosphorylation were examined. Protein tyrosine phosphorylation preceded PLD activation, and a good correlation was also shown in the effect of genistein between these two events—inhibited phosphorylation and decreased PLD activation. Moreover, in the presence of vananade, there was potentiation in both protein tyrosine phosphorylation and PLD activation induced by H2O2, suggesting involvement of PTK in H2O2-induced PLD activation in vascular endothelial cells. The effect of H2O2 on activation of PLD has been studied in other cell types such as pheochromocytoma PC12 cells, lymphocytic leukemic cells, and Swiss 3T3 fibroblasts. In PC12 cells, it has been shown that exposure to H2O2 markedly stimulates PLD activity in a time- and concentration-dependent manner. Pretreatment with PTK inhibitors but not PKC inhibitors diminished PLD activation (Ito et al., 1997b). There is a synergistic increase in H2O2-mediated PLD activity in PC12 cells by the addition of vanadate. Such stimulation of PLD activity by H2O2 is not observed in cell lysates. Depletion of extracellular calcium abolishes H2O2-induced PLD activation, which is not consistent with the case of vascular endothelial cells (Natarajan et al., 1996). A calcium-dependent PTK, Pyk2 has been

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suggested as an important factor for PLD activation by oxidant. Although these observations strongly suggest that tyrosine phosphorylation stimulates PLD activity directly or indirectly, the precise mechanism remains unclear. In another study with PC12 cells overexpressing PLD2, H2O2-induced stimulation of PLD is sensitive to PKC inhibitors but less sensitive to PTK inhibitors, indicating that PLD activation by H2O2 is largely dependent on PKC and calcium and minimally dependent on PTK (Oh et al., 2000). The different profile of sensitivity to the protein kinase inhibitors might be due to difference in strains used or experimental conditions. The sensitive response to both PKC and PTK inhibitors in H2O2-induced PLD activation is also found in Swiss 3T3 fibroblasts (Min et al., 1998) and A431 carcinoma cells that express PLD1 and PLD2 (Min et al., 2001a). H2O2 induces tyrosine phosphorylation of both PLD isozymes in A431 cells. Recent work has shown that in PC12/PC2 cells, peroxyvanadate causes direct phosphorylation on tyrosine residues of PLD2 and an increase in PLD activation, but there is a lack of correlation between protein phosphorylation and activation of PLD1 in terms of time course and dose dependency (Mehta et al., 2003). Thus, despite abundant data suggesting possible participation of tyrosine phosphorylation in PLD activation induced by oxidant, which type of PTK is involved and how it functions in the process are still in dispute. Src has been thought to act as a major PTK for phosphorylation by studies using its inhibitors. Recent study has demonstrated the PLD2 is associated with Src and Pyk2 in PC12 cells (Banno et al., 2005), but there is no evidence to indicate direct phosphorylation by Src of PLD2. More recently, coexpression experiments of v-Src and PLD1 in COS-7 cells showed increased activity and marked tyrosine phosphorylation of PLD1. However, the results do not support the implication of tyrosine phosphorylation in the activation of PLD1, though the association of the two proteins is important for the activation of PLD1 (Ho et al., 2005). Another report has provided evidence for tyrosine phosphorylation of PLD1 by Src via direct association in A431 cells (Ahn et al., 2003).

3.4 Mitogen-Activated Protein (MAP) Kinase The MAP kinase family of serine/threonine kinases comprises three main subgroups including the extracellular signal-regulated kinase (ERK), the p38 MAP kinase (p38MAPK), and the c-jun N-terminal kinase, also known as stress-activated protein kinase. These MAP kinases are activated by dual phosphorylation on closely located threonine and tyrosine residues. Several studies have shown that MAP kinase is implicated in activation of PLD induced by oxidant- (Ito et al., 1998; Banno et al., 2001; Natarajan et al., 2001; Min et al., 2002) and receptor-mediated agonists (Muthalif et al., 2000; Watanabe et al., 2004b; Li and Malik, 2005; Paruch et al., 2006). However, most of evidence is based on the fact that inhibitors of MAP kinase inhibit activation of PLD. It was first demonstrated that, in PC12 cells exposed to H2O2, stimulation of PLD activity and phosphorylation of ERK are dose dependently inhibited by a specific inhibitor for a MAP kinase-ERK kinase (MEK), PD098059 (Ito et al., 1998). In contrast, carbachol-mediated PLD activation is not affected by the inhibitor, whereas ERK phosphorylation is blocked. Furthermore, when PC12 cells are treated with the p38MAPK inhibitor, SB203580, prior to stimulation by H2O2, PLD activation is reduced in a time- and dose-dependent manner (Banno et al., 2001). A combination of these two MAP kinase inhibitors causes much greater inhibition of PLD activation. These observations have tempted us speculate that ERK and p38MAPK are involved in PLD activation induced by H2O2 exposure. Implication of p38MAPK in peroxyvanadate-induced PLD activation has also been observed in vascular endothelial cells (Natarajan et al., 2001). The p38MAPK inhibitor or transfection of its dominant negative mutant diminishes the PLD activation. Immunoprecipitation experiments reveal that PLD1 and PLD2 are associated with p38MAPK. Both PLD isozymes can be phosphorylated by p38MAPK in vitro, and peroxyvanadate enhances their phosphorylation. However, phosphorylation of PLD by p38MAPK causes no significant effect on PLD activity. In addition to involvement of ERK and p38MAPK in oxidant-induced PLD activation, there are several lines of evidence for the role of these MAP kinases in receptor-mediated PLD regulation; ERK in vascular smooth muscle cells (nor-epinephrine) (Muthalif et al., 2000), PC12 cells (NGF) (Watanabe et al., 2004a), cerebellar granule neurons (adhesion molecule L1) (Watanabe et al., 2004b), and p38MAPK in vascular

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smooth muscle cells (angiotensin II) (Li and Malik, 2005). The results obtained from these experiments in various cell types indicate that ERK and p38MAPK act directly or indirectly upstream of agonist-induced PLD activation. Recent study has provided potential evidence of ERK/PLD2 coupling induced by a chemoattractant peptide fMLP in HL60 cells or HEK cells stably expressing fMLP receptors (Paruch et al., 2006). ERK inhibition by the MEK inhibitor U0126 abolishes fMLP-mediated stimulation of PLD activity. Transfection of a constitutively active MEK1 results in potentiated activation of PLD2. PLD2 coimmunoprecipitates with ERK and is phosphorylated on the MAP kinase consensus sequence S/T-P in fMLP-stimulated cells. Moreover, incubation of plasma membranes with a recombinant active ERK in the presence of ATP causes a rapid and transient PLD activation. Direct phosphorylation of PLD2 is also confirmed in the cell-free system in the presence of ERK and ATP. It has, on the other hand, been argued whether PLD activation occurs downstream or upstream of ERK, but the relationship of ERK with the PLD signaling is most likely dependent on the agonist and the cell type.

4

Functions of Phospholipase D

Much work on functions of PLD has revealed that PA generated by this enzyme is a multifunctional lipid (McDermott et al., 2004; Jenkins and Frohman, 2005; Cazzolli et al., 2006). It is widely acknowledged that PA as a bioactive lipid second messenger is linked to diverse events of intracellular signaling, as depicted in > Figure 15-1. PA serves as an activating factor of various enzymes, including PIP5-kinase, sphingosine kinase1, mTOR (mammalian target of rapamycin) kinase, Raf-1 kinase, and NOX (NADPH oxidase). In contrast, PA binds specifically and directly to the protein phosphatase1 catalytic subunit, leading to inhibition of its enzyme activity (Jones and Hannun, 2002). PA is also important for the production of

. Figure 15-1 PLD/PA signaling PC, phosphatidylcholine; ERK, extracellular signal-regulated kinase; PKC, protein kinase C; PI (4,5)P2, phosphatidylinositol(4,5)bisphosphate; PLD, phospholipase D; DG, 1,2-diacylglycerol; DGK, diacylglycerol kinase; PAP, phosphatidic acid phosphatase; PLA, phospholipase A; LPA, lysophosphatidic acid; PI3K, phosphatidylinositol 3-kinase; PP-1, protein phosphatase-1; SPHK, sphingosine kinase; mTOR, mammalian target of rapamycin; S6K1, ribosomal p70S6 kinase 1; PIP5K, PI4P 5-kinase; PDE; phosphodiesterase

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other lipid-signaling mediators; DAG generated from PA by PA phosphatase and LPA generated from PA by phospholipase A2. Additionally, PA plays a biophysical role; the conversion of PC to PA by PLD causes a perturbation in the localized membrane lipid domain, and PA can also act as a potent fusogenic lipid. Both effects are required for the vesicle transport (McDermott et al., 2004; Jenkins et al., 2005). Thus, as a second messenger and also as a biophysical membrane perturbant, PA is implicated in a variety of cellular functions, e.g., cytoskeletal rearrangement, vesicular trafficking, exocytosis, endocytosis, mitogenesis, survival, lipid droplet formation, and melanogenesis. Here, some of the physiological events elicited by this pleiotropic lipid are described. Insulin-stimulated uptake of glucose is mediated by the Glut4 glucose transporter. Upon stimulation by insulin, cytosolic vesicles containing Glut4 translocate to the plasma membrane for fusion by which the transporter is exposed to the extracellular milieu. The Glut4 exocytosis is thought to be a crucial step in the whole Glut4 translocation process that controls the rate of glucose uptake. PA has been indicated to function as an anchor for many proteins including the SNARE (soluble N-ethylmaleimide-sensitive factor attachment protein receptor) complex (Huang and Frohman, 2003). Interaction of the vesicles with the plasma membrane is mediated by SNAREs located on secretory vesicles (v-SNARE) and on the plasma membrane (t-SNARE). In insulin-stimulated cells, these proteins form a stable complex leading to the vesicle docking to the plasma membrane before fusion. PA also plays an important role as a fusogenic lipid in the fusion of the Glut4-containing vesicles (Huang et al., 2005). Membrane fusion is a remodeling process requiring energy to overcome the strong hydrophobic effect of the inside of lipid bilayer. Coneshaped PA with its small negative charge induces negative curvature at the inward membrane curve. Thus, production of PA by PLD is a key event in the late fusion process of Glut4-containing vesicles. In fact, it has recently been demonstrated that, in insulin-stimulated 3T3-L1 adipocytes, the increase of PLD1 activity by its overexpression enhances Glut4 translocation and glucose uptake, whereas decreased PLD1 activity by transfection of its siRNA or catalytically inactive mutant causes inhibition of these events. More recently, SNARE-dependent membrane fusion has been exhibited in a reconstituted system (Vicogne et al., 2006). Addition of PA to Stx/SNAP23 vesicles results in marked enhancement of the fusion rate, whereas its addition to VAM2 vesicles is inhibitory. Interestingly, addition of PI(4,5)P2 to Stx/SNAP23 vesicles is inhibitory for fusion, but its addition to VAM2 vesicles is stimulatory. These observations provide compelling evidence that concerted distribution of these acidic phospholipids is beneficial for complete fusion. An intriguing finding has been presented to suggest involvement of PLD1 in insulin-induced lipid droplet formation (Andersson et al., 2006). Lipid droplets, which consist of a core of neutral lipids surrounded by a monolayer of phospholipids, increase in size by fusion of small droplets. Insulin stimulates the formation of lipid droplets. Increased activity of PLD by ectopic expression of PLD1, but not PLD2, enhances the formation of lipid droplets, whereas knockdown of PLD1 with siRNA inhibits droplet formation, thereby suggesting a pivotal role of PLD1 in this event. In the cell-free system, addition of PA promotes the formation of droplets. It can be assumed that PA may recruit other proteins involved in the process or may function as a biophysical perturbant of membrane lipid bilayers for fusion. Another crucial factor in droplet formation through PLD signaling is ERK2. Overexpression and downregulation with siRNA of ERK2 reveal its essential involvement. ERK2 increases phosphorylation of dynein, which is important for the interaction between dynein and ADRP-containing droplets. Although it is highly conceivable that both PLD1 and ERK2 play essential roles in the insulin-stimulated accumulation of lipid droplets, the exact relationship between these two enzymes-involving pathways remains unclear. Of interest is a unique functional role of PLD in melanogenesis. The melanogenic process is a highly regulated and complex event occurring in melanocytes and melanoma cells, and is initiated by activation of tyrosinase, the rate-limiting enzyme. Ohguchi et al. (2004) have come across the interesting finding that pigmented B16 melanoma cells show no significant expression of PLD1 as examined by immunofluorescent staining, whereas nonpigmented cells abundantly express PLD1. It has also been demonstrated that PLD activity decreases and tyrosinase activity increases during the progression of melanogenesis induced by a-MSH. The melanin content and the tyrosinase activity and expression are markedly reduced in PLD1-overexpressed cell. The downregulation of PLD1 with its siRNA exhibits the opposite effects, i.e., increases in the melanin content and the activity and expression (mRNA and protein) of tyrosinase. Furthermore, phosphorylation of p70S6K is diminished in PLD1-knockdown cells, and rapamycin, a

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potent inhibitor of mTOR that is an upstream effector of p70S6K, induces a marked melanogenesis (Ohguchi et al., 2005). These observations constitute strong evidence for the PLD1’s negative regulation in the melanogenic process through mTOR/p70S6K signaling.

5

Role of Phospholipase D Signaling in Neural Cells

PLD isoforms are present in neurons and glial cells (astrocytes and oligodendrocytes), as well as glioma, neuroblastoma, astorocytoma, and pheochromocytoma cell lines, and stimulation of the cells with various agonists including neurotrasmitters, hormones, oxidants, and growth factors induces activation of PLD to various extents depending on the cell type and the agonist (Klein, 2005). Most of the PLD-linked cellular functions as described above can also be observed in neural cells. This section considers evidence for implication of PLD activation in neurotransmitter release, differentiation (neurite outgrowth), survival (antiapoptosis), and neurodegeneration.

5.1 Neurotransmitter Release The role of PLD in secretion has been shown mainly in neuroendocrine cells, but has also been studied in mast cells, adipocytes, and pancreaticbcells. Many homologies can be seen in the PLD-dependent exocytotic machinery between these cells and neuronal cells. In neuroendocrine chromaffin cells, there is a causal relationship between activation of PLD1 and secretion of catecholamine (Vitale et al., 2001). Moreover, overexpression of wild-type PLD1 in PC12 cells increases secretion of growth hormone as a parameter of exocytosis, whereas the catalytically inactive PLD1 is inhibitory. These findings provide the first direct evidence for a crucial role of PLD in the secretory machinery in neuroendocrine cells. Such an important role of PLD1 in exocytosis has also been demonstrated in acetylcholine (ACh) release at synapses (Humeau et al., 2001). Microinjection of catalytically inactive PLD1 into a presynaptic cholinergic neuron in the Aplysia buccal ganglion resulted in a profound inhibition of ACh release, most likely by decreasing the number of functional presynaptic releasing sites. Upon stimulation with agonists, the secretory vesicles underwent rapid fusion with the plasma membrane leading to exocytosis. During the fusion process, several events occur such as docking the vesicles to the plasma membrane, priming, and fusion pore formation. By analogy with the function proposed in neuroendocrine cells, PLD1is assumed to induce the biophysical modification of the membrane lipid bilayers at the fusion sites of presynaptic plasma membrane. Despite the importance of PLD1 activation in generating PA in the SNARE-mediated fusion process, the orientation of the factors required for activation of PLD1 has not been defined. Recent study has established that SCAMP2 (secretory carrier membrane protein2) localized on the plasma membrane functions as a scaffold on which Arf6 is activated by ARNO, a guanine nucleotide exchange factor, and PLD1 is then activated by Arf6-GTP and PI(4,5)P2 (Liu et al., 2005). Considering that PI(4,5))P2 is produced by PIP5-kinase, which is activated by Arf6 and PA, a concerted cross talk between PLD1, PIP5-kinase, ARNO, Arf6, PA, and PIP2 is thought to regulate the formation of fusion pores.

5.2 Differentiation (Neurite Outgrowth) Several lines of evidence have indicated that PLD is associated with differentiation in various cell types, such as myeloid cell lines (HL60 and NB4), keratinocytes, and neuronal cells (Nakashima and Nozawa, 1999). The initial finding was the increased mRNA expression of PLD isozymes (PLD1a, PLD1b, PLD2) during granulocytic differentiation induced by dibutyryl cyclic AMP (dbcAMP) or ATRA in HL60 cells (Nakashima et al., 1998). It has also been reported that PLD activity and expression of PLD1 and PLD2 are increased in relation to maturation and differentiation in human leukemic myeloid cells ex vivo (El Marjou et al., 2000). C6 glioma cells are induced to express astrocytic phenotypes by the treatments that increase intracellular cyclic AMP level. During differentiation by dbcAMP, differential mRNA expression of

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PLD isozymes and increased PLD activity have been observed (Yoshimura et al., 1996). Most of the early studies have been done in PC12 cells and suggest a possible role for PLD in neurite outgrowth. First, it was shown that expression of PLD1 mRNA increases during formation of neurites induced by dbcAMP or NGF (Hayakawa et al., 1999). Upregulation of PLD1 as well as PKC (a and bII) protein expression and the increased association of these two proteins implicate PKC-dependent PLD activity in NGF-mediated differentiation of PC12 cells (Min et al., 2001b). It has also been exhibited that increased expression of wild-type PLD2 causes enhanced elongation of neurites induced by NGF stimulation or transient expression of constitutively active MEK in PC12 cells and furthermore that the MEK inhibitor PD98059 prevents both NGF-induced PLD2 activation and PLD2-enhanced neurite outgrowth (Watanabe et al., 2004b). These observations strongly suggest that PLD2 activation occurring downstream of ERK pathway is a critical step for neuronal differentiation. Recent study has shown an induction of neurite outgrowth by lysophosphatidylcholine (LPC) through activation of PLD2. Studies with PC12 cells have been extended to those with other neuronal cells. In immortalized hippocampal stem cells (HiB5), stimulation by PDGF, a neurogenic factor, results in a rapid and transient activation of PLD and phospholipase C (PLC), and inhibition of either of the enzymes reduces neurite outgrowth, indicating that both PLD and PLC may contribute to neural differentiation by PDGF (Sung et al., 2001). Basic FGF induces neurite outgrowth concurrent with PLD activation in another hippocampal cell line (H19–7) (Oh et al., 2007). Overexpression of active PLD1 dramatically increases bFGF-induced neurite outgrowth and increases PLD activity, whereas transfection of its inactive mutant shows the opposite effect. It is to be noted that bFGF-mediated PLD1 activation via PLC-g and neurite formation does not require the ERK pathway. In the in vivo model of hippocampal mossy fiber outgrowth, the expression levels of both PLD1 and PLD2 increase along the path of mossy fiber sprouting induced by kainic acid injection into the amygdala (Zhang et al., 2004). Moreover, it is of interest from the pathophysiological aspect that PLD1 enhances the release of tissue plasminogen activator (tPA) leading to hippocampal mossy fiber outgrowth, because epileptic seizures are modulated by tPA that promotes a proteolytic action required for neurite extension (Zhang et al., 2005). Taken together, the evidences described here strongly support the proposal that PLD activation plays an important part in neuronal differentiation, but further experiments are required to understand the precise mechanisms for the downstream events elicited by PLD signaling.

5.3 Survival (Antiapoptosis) Apoptosis is a critical feature of embryogenesis, cell growth, and differentiation in multicellular organisms. In the developing nervous system, neurons are generated in excess and compete for limited survival factors, and a large member of cells are eliminated by the apoptotic process. Additionally, mammalian cells undergo apoptosis by exposure to multiple diverse stimuli, including TNFa, ionizing radiation, chemotherapeutic agents, and oxidative stress. In order to overcome or prevent apoptotic death mammalian cells are endowed with the antiapoptotic (survival) machinery that functions through various signal transduction pathways, including phospholipid signaling. The PI3K/Akt pathway is a typical survival signaling system. The antiapoptotic role of this pathway through PLD has also been demonstrated in CHO cells treated with actinomycinD (ActD) (Yamada et al., 2004). Overexpression of either PLD1 or PLD2 protects cells from ActD-induced apoptosis. In the early phase of the apoptotic process, ActD induces an increase in PLD activity concurrent with activation of the PI3K/Akt/p70S6K pathway. More recently, elevated expression of either PLD1 or PLD2 has been shown to prevent etoposide-induced apoptosis by inhibiting expression of the tumor suppressors, early growth response-1 (Egr-1), and PTEN, providing evidence for a novel mechanism for counteracting apoptosis (Kim et al., 2006). As described above, oxidative stress (H2O2) induces PLD activation in PC12 cells, but its physiological significance has been unknown (Ito et al., 1997b). However, the finding that the activation of PLD precedes the appearance of apoptotic features has led to a proposal suggesting PLD’s role in survival signaling. More direct data to support the proposal have been found by transfection of the active PLD2 and its inactive mutant (Lee et al., 2000). H2O2-induced apoptosis is suppressed in PLD2-overexpressed PC12 cells,

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whereas it is enhanced in the inactive mutant-transfected cells. Further, PC12 cells undergo apoptosis following exposure to hypoxic shock by accumulating ceramide produced via sphingomyelinase (Yoshimura et al., 1998), but overexpression of active PLD2 significantly inhibits apoptotic cell death because of hypoxia (Yamakawa et al., 2000). Despite this supportive evidence implicating PLD as a critical regulator of survival signaling, its molecular mechanisms have been unclear. Recent study has shown that PLD2 is required for H2O2-enhanced association of Src with Pyk2 leading to full activation of pyk2 and that oxidant-induced activation of PI3K/Akt is reduced by transfection of catalytically inactive PLD2 mutant in PC12 cells (Banno et al., 2005). It can thus be assumed that PLD2 activation induces Src-dependent activation of Pyk2 by promoting the complex formation between Pyk2 and activated Src, thereby resulting in activation of the survival signaling pathway PI3K/Akt/p70S6K. An attractive novel mechanism underlying PLD-dependent survival signaling has been proposed by implicating mTOR as a direct target for PA generated by PLD (Fang et al., 2001). In fact, PLD2, but not PLD1, has recently been shown to form a functional complex with mTOR/raptor (Ha et al., 2006). mTOR is a serine/threonine kinase that regulates cell growth and proliferation through the downstream effectors, p70S6K, and eukaryotic translation initiation factor 4E binding protein1 (4E-BP1). Many lines of evidence indicate that mTOR is activated via PI3K/Akt signaling. The emergence of the alternative PLD/mTOR pathway gives rise to a new insight into the role of PLD in survival signaling (Foster, 2007). In HEK 293 cells, overexpression of wild-type PLD1 causes an increase in p70S6K activity, whereas its catalytically inactive mutant has a negative effect on p70S6K activity (Fang et al., 2003). PLD1 siRNA also inhibits 4E-BP1 and p70S6K1 phosphorylation. In human breast cancer cells, it has been reported that cells with elevated PLD activity undergo mTOR-dependent survival signaling and that the cells with very low PLD activity are dependent on the PI3K-survival pathway (Chen et al., 2005). In addition to suppression of apoptosis induced by serum withdrawal, high PLD activity also enhances cell migration and invasion, postulating a dual role of the PLD signaling in tumorigenesis (Zheng et al., 2006). An interesting study has revealed that mutant p53 is stabilized by elevated PLD activity and participates in survival signaling through PLD in MDA-MB-231 cells exposed to serum deprivation (Hui et al., 2006; Lehman et al., 2007). Transfection of siRNA for PLD2 reduces p53 expression and suppresses the phosphorylation of MAP kinase, but not MEK, and the p53 expression is insensitive to rapamycin, a specific inhibitor for mTOR. These findings strongly support that the PLD-dependent expression of p53 is mediated by MAP kinase, but not mTOR. As described here, an established correlation between PLD and mTOR has been well documented. However, a novel mechanistic potential of PLD in p70S6K activation has only recently been postulated (Lehman et al., 2007). Although overexpression of active PLD2 results in elevated activities of PLD and p70S6K in COS-7 cells, overexpression of its inactive mutant has the opposite effects. Moreover, either rapamycin treatment or knockdown of mTOR does not affect the PLD-dependent stimulation of p70S6K activity, thus implying a direct binding of PA to and activation of p70S6K, which is independent of mTOR. Although the PLD-mediated p70S6K activation, either dependent on or independent of mTOR, may play a protective or survival role of PLD in the nervous system, more work is needed to determine the molecular targets of this signaling pathway.

5.4 Neurodegeneration The data discussed above implicate PLD signaling in the antiapoptotic process and suggest its beneficial role for neurodegenerative disorders. Kanfer et al. (1986) have proposed that PC present in neuronal cell membranes could be a precursor of neurotransmitter ACh that is produced from choline by choline acetyltransferase. As choline is the product of PLD, they expected reduced activity of PLD in brain tissue samples from Alzheimer’s disease (AD). Indeed, there was a marked reduction in the choline acetyltransferase and PLD activities. This highlights another important function of PLD as a key enzyme in ACh synthesis, in addition to many other physiological functions (Klein, 2005). Recent study has shown a close correlation between the expression and activity of PLD2 and ACh synthesis, as inferred by transfection of PLD2 or its antisense oligonucleotide in cholinergic SN56 cells (Zhao et al., 2001).

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It has been established that b-amyloid (bA), which is produced by g-secretase cleavage of bA precursor protein (bAPP), is a critical factor in the progress of AD. Presenilin1, as a major component of g-secretase, binds to and recruits PLD1 to TGN (trans-Golgi network). Of particular interest, ectopic expression of wild-type PLD1 reduces bA production, whereas downregulation by transfection of its siRNA enhances bA production in N2a neuroblastoma cells (Cai et al., 2006a). Moreover, the active PLD1 overexpression increases bAPP trafficking and the inactive PLD1 expression does not. It appears, therefore, to be plausible that the suppression of bA generation by PLD1 may result from impaired trafficking of bAPP. However, from the data showing that overexpression and downregulation of PLD1 decreases and increases the g-secretase activity, respectively, it is most likely to consider that the PLD1-mediated reduction of bA production is principally due to g-secretase inhibition, through the dissociation of the PS1/g–secretase complex. Further related work has reported that the elevated PLD1 activity in familial AD mutant neurons rescues aberrant bAPP trafficking and impaired neurite outgrowth (Cai et al., 2006b). These findings indicate that PLD1 could be of preventive or therapeutic benefit with AD as a potential target.

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Glutamate Receptors: NMDA and Delta Receptors

M. Yuzaki

1

Introduction – Why NMDA and Delta Receptors? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 316

2 2.1 2.2 2.3 2.3.1 2.3.2 2.3.3 2.3.4 2.4 2.4.1 2.4.2

NMDA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 316 Structure and Basic Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 316 Genes and Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 319 Trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 320 ER Exit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 320 Transport Along Dendrites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 320 Exocytosis/Endocytosis at Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 320 Synaptic Versus Extrasynaptic NMDA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 321 Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 321 Functional Diversities of NR1/NR2 Complexes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 321 Functions of NR3-Containing NMDA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 322

3 3.1 3.2 3.3 3.3.1 3.3.2 3.4 3.4.1 3.4.2

Delta Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 322 Structure and Basic Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 322 Genes and Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 323 Trafficking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 324 ER Exit and Transport along Dendrites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 324 Exocytosis/Endocytosis at Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 324 Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 324 Phenotypes of Glurd2-Null Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 324 Mechanisms Clarified by the “Transgenic Rescue” Approach . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 326

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Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 327

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2009 Springer ScienceþBusiness Media, LLC.

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Glutamate receptors: NMDA and delta receptors

Abstract: N-methyl-D-aspartate (NMDA) receptors and delta (d) receptors are members of the ionotropic glutamate receptor (iGluR) family, along with a-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors and kainate receptors. Unlike AMPA receptors, both NMDA and d receptors do not contribute to normal fast neurotransmission. Instead, the activity of these receptors regulates AMPA receptor trafficking and synaptic plasticity. The traditional function of NMDA receptors is achieved through their Ca2+ permeable channels. However, it has become increasingly clear that intracellular proteins that bind to the C-termini of NR2 subunits, and possibly those of NR3 subunits, play crucial roles in NMDA receptor trafficking and intracellular signaling, leading to specific gene expression. Similarly, recent studies have indicated that the d2 glutamate receptor does not function as a channel, but as a nonionotropic receptor that regulates intracellular signaling through its C-terminal interaction. Another feature common to both NMDA and d2 receptors is their ability to bind to D-serine and glycine; these molecules are released from glial cells as “gliotransmitters” and regulate synaptic plasticity. Therefore, although the amino acid sequence similarity between the two receptor families is rather low, characterization of the signaling mechanisms underlying each family should add to the knowledge of each family, facilitating a better understanding of learning and memory. List of Abbreviations: AMPA, a-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid; BDNF, brainderived neurotrophic factor; CaN, calcineurin; CREB, cyclic AMP responsive element-binding protein; ER, endoplasmic reticulum; ERK, extracellular signal-regulated kinase; iGluR, ionotropic glutamate receptor; JNK, c-Jun NH2-terminal kinase; LBD, ligand-binding domain; LTD, long-term depression; LTP, longterm potentiation; MAPK, mitogen-activated protein kinase; NMDA, N-methyl-D-aspartate; NTD, Nterminal domain; PF, parallel fiber; PKC, protein kinase C; PSD, postsynaptic density; SynGAP, synaptic Ras GTPase activating protein; TM1, transmembrane domain 1

1

Introduction – Why NMDA and Delta Receptors?

Ionotropic glutamate receptors (iGluR) consist of a-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors, kainate receptors, N-methyl-D-aspartate (NMDA) receptors, and delta (d) receptors. Because of their essential roles in many neurodevelopmental, neurophysiological, and neuropathological processes, numerous papers, including excellent reviews, have been published in the field of NMDA receptors in the past decades. In contrast, the d receptors are the least characterized subfamily of iGluRs and had been regarded as “orphan receptors” until very recently. Although the delta family shares a fulllength amino acid sequence similarity of only 20–30% with the NMDA receptor family, these two families have one thing in common – they both bind to glycine and D-serine (Furukawa et al., 2003; Naur et al., 2007). In addition, both receptors do not contribute to fast excitatory neurotransmission, which is mainly mediated by AMPA receptors, but rather serve to regulate synaptic plasticity and synaptogenesis during development and in mature brain. Because several aspects of each receptor family have been reviewed with original citations (Yuzaki, 2003; Lau et al., 2007; Paoletti et al., 2007; Gereau and Swanson, 2008), relatively new findings discovered in recent years would be focused, without referring to some original reports.

2

NMDA Receptors

2.1 Structure and Basic Properties NMDA receptors consist of three different subfamilies: NR1, NR2 (NR2A, NR2B, NR2C and NR2D), and NR3 (NR3B and NR3B). Like other iGluR families, the extracellular region includes the N-terminal domain (NTD), the sequence of which is similar to that of bacterial periplasmic protein leucine/isoleucine/valinebinding protein, and the ligand-binding domain (LBD), which is evolutionally conserved from another bacterial periplasmic protein lysine/arginine/ornithine-binding protein. The LBD consists of two segments,

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S1 and S2, separated by a transmembrane domain 1 (TM1), an ion channel-forming re-entrant loop segment (TM2), and TM3 (> Figure 16-1). The cytoplasmic C-terminus region following TM4 binds to many intracellular signaling and anchoring molecules (> Figure 16-1). One of the major features of NMDA receptor channels – their high permeability to Ca2+ and voltagedependent blockade by Mg2+ (> Figure 16-2, > Table 16-1) – is achieved by two asparagine (N) residues located at TM2 (> Figure 16-1). The first N site is homologous to the glutamine/arginine (Q/R) site, which regulates the Ca2+ permeability of AMPA and kainate receptors. In NR3 subunits, glycine (G) occupies the corresponding site; thus, NMDA receptors containing NR3 subunits display reduced Ca2+ permeability (Perez-Otano et al., 2001; Matsuda et al., 2003), but the degree of blockade by Mg2+ is similar to that in NMDA receptors without NR3 subunits (Nishi et al., 2001; Yamakura et al., 2005). Zn2+ binds to the NTD of NR2 subunits and inhibits NMDA currents with an IC50s (mM) of 0.02 (NR2A), 2 (NR2B), 20 (NR2C), and 10 (NR2D) (Paoletti et al., 2007). H+ binds to the NTD of NR1 near the N1 region and post-TM3 regions of NR1 and NR2 subunits and mediates the inhibition of NMDA currents under low pH conditions (> Figure 16-1) (Low et al., 2003). The NTD of NR1 is also required for homodimerization of NR1 subunits. D-serine and glycine bind to the purified LBD of NR1 with KD values of 7.02 and 26.4 mM, respectively (Furukawa et al., 2003). D-serine and glycine bind much more strongly to the purified LBD of NR3A, with

. Figure 16-1 Membrane topology and associated intracellular molecules of NMDA receptors. The putative ligand-binding domain (LBD), formed by the S1 and the S2 regions, is separated by transmembrane (TM) domains 1 through 3. The ligand-binding regions for typical ligands are indicated by arrows. The most N-terminal domain (NTD) is also indicated. The ion selectivity filter in the TM2 is encoded by asparagine (N) for NR1 and NR2 and by glycine (G) for NR3. Intracellular molecules that bind to the C terminus of NMDA receptors are indicated with their proposed functions. The C terminus of NR1 consists of C0, C1, and C2/C20 cassettes. PDZ proteins bind to the C20 or SXV motifs at the end of the C-terminus of NR1 or NR2, respectively. The C0 cassette is involved in the desensitization of NMDA receptor currents. NR2B binding to CaMKIIa via two regions (proline-rich region, P; central region, C) can produce the sustained activity of CaMKIIa. Proximal tyrosine residues (Y) of NR1 and NR2 are used for sorting to late endosomes, while the LL motif of NR2A and the YEKL motif of NR2B are recognized by AP-2 and sorted to recycling endosomes

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. Figure 16-2 Function of NMDA receptors. The function of NMDA receptors is achieved through their Ca2+ permeable channels, which are blocked by Mg2+ in a voltage-dependent manner. Large increases in Ca2+ levels preferentially activate CaMKIIa, leading to long-term potentiation (LTP), whereas small increases activate calcineurin (CaN) and long-term depression (LTD). In addition to such ionotropic actions, the functions of NMDA receptors are regulated by nonionotropic actions via molecules that bind to the C-terminus of NR2, leading to gene expression changes

. Table 16-1 Channel properties formed by NR1 and various NR2 subunits NR2 subunit NR2A NR2B NR2C NR2D

Single channel conductance (pS)a 50, 40 50, 40 35, 22 35, 16

Sensitivity to extracellular Mg2+(IC50; mM)b ++(20) ++(20) +(80) +(80)

Kinetics (deactivation time constants; msec)c Fast (33–70, 247–350) Slow (71, 538) Slow (260–376) Slowest (45, 4408)

a

Cull-Candy et al. (2004) Paoletti et al. (2007) c Dingledine et al. (1999) b

KD values of 643 and 40 nM, respectively (Yao et al., 2006). Considering the concentration of these amino acids in the brain, NR3A subunits are likely saturated by glycine or D-serine in vivo. In contrast, although binding assays on the purified LBD of NR2 have not been performed, the LBD of NR2 provides a binding site for glutamate; glutamate evokes a current in cells expressing NR2B with an EC50 of 1.5 mM (Laube et al., 1997). Although the stoichiometry of native NMDA receptors has not been established unequivocally, NMDA receptors are most likely tetramers consisting of two NR1 and two NR2 subunits of the same or different subtypes. Immunoprecipitation analysis of hippocampal lysates indicates that each NR1/NR2A, NR1/ NR2B, or NR1/NR2A/NR2B complex constitutes approximately one-third of the total NMDA receptor population (Al-Hallaq et al., 2007). The properties of tandem-linked NMDA receptors (a fusion of NR1NR1, NR2-NR2, or NR1-NR2 subunits) suggest a dimer of homodimers (NR1-NR1 + NR2-NR2), as opposed to a dimer of heterodimers (NR1-NR2 + NR1-NR2) (Schorge et al., 2003; Papadakis et al., 2004).

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The stoichiometry of NMDA receptors containing the NR3 subunit is less clear. In contrast to the assembly mechanism underlying NR1/NR2 complexes, NR1/NR2/NR3 complexes have been recently shown to be composed of a dimer of heterodimers (NR1-NR2 + NR1-NR3) (Schuler et al., 2008). In response to the sustained application of ligands, NMDA currents are gradually reduced by a phenomenon called desensitization. In AMPA receptors, the rearrangement of the LBD dimer-dimer interface is coupled to its strikingly rapid and thorough desensitization, with a time constant of a few milliseconds. In contrast, NMDA receptors desensitize much more slowly (> Table 16-1). It is unclear whether the desensitization of NMDA receptors is also affected by the rearrangement of LBD dimers. However, at least three kinds of mechanisms of NMDA receptor desensitization are known (Dingledine et al., 1999). Glycine-dependent desensitization is caused by an allosteric reduction of glycine binding to the LBD of NR1 by glutamate binding to the LBD of NR2. A glycine-independent and Ca2+-dependent form of desensitization is mediated by the direct binding of Ca2+-calmodulin to the C0 and C1 region (> Figure 16-1). Ca2+-calmodulin-dependent protein kinase II a (CaMKIIa) and a-actinin also bind to the C0 region and modify the Ca2+-dependent desensitization process (Leonard et al., 2002). The intracellular loop between TM2 and TM3 of the NR2 subunit is also speculated to interact with the C0 region of the NR1 subunit and to modulate Ca2+-dependent desensitization (Vissel et al., 2002). Finally, a glycineinsensitive and Ca2+-insensitive form of desensitization has been observed in NR1-NR2A and NR1-NR2B receptors but not in those with NR2C and NR2D. Although the precise mechanisms are unclear, this form of desensitization is mediated by the region preceding the TM1 of NR2 (Thomas et al., 2006a) and are modulated by protein kinase C activities (Jackson et al., 2006).

2.2 Genes and Expression NMDA receptors are encoded by a total of seven genes – one for NR1, four for the NR2 subunits (NR2A, NR2B, NR2C and NR2D), and two for the NR3 subunits (NR3A and NR3B). The NR1 subunit exists in eight different variants generated by alternative splicing at three sites within the protein, one in the N terminus (N1 cassette) and two in the C-terminus (C1 and C2 cassettes); when the C2 cassette is absent, the separate C20 cassette is used instead (> Figure 16-1). NR1 mRNA is expressed ubiquitously in virtually all neurons from as early as embryonic day (E) 13 through adulthood in mice (Gereau and Swanson, 2008). NR2A mRNA is a major form in postnatal brain, while NR2B mRNA is predominant in embryonic brain, indicating specific roles of each subunit (see > Section 2.3.4). In adult, NR2B mRNA is still expressed but is restricted to forebrain regions. In the cerebellum, NR2B mRNA is replaced by NR2C mRNA during postnatal development. NR2D mRNA is mainly expressed in the diencephalon and the brainstem at embryonic and neonatal stages. NR3A is expressed in wide regions of the brain during the first postnatal week, but its expression decreases to a low level in adult brain, except for in restricted regions like the olfactory bulb (Wong et al., 2002) and cerebellar interneurons (Fukaya et al., 2005b). High levels of NR3B mRNA and protein are expressed in motoneurons in the brainstem (trigeminal motor, facial, and ambiguous nuclei) and upper spinal cord (Nishi et al., 2001), but low levels of NR3B protein are widely expressed in the forebrain and the cerebellum (Wee et al., 2008). The mechanism by which NR2B is replaced by NR2C in cerebellar granule cells has been recently characterized and proposed to occur as follows (Nakanishi et al., 2006): when granule cells are located in the external granule layer, they are depolarized to about 25 mV by ambient glutamate, and Ca2+ influx through voltage-gated Ca2+ channels activates CaMKIIa, leading to the expression of brain-derived neurotrophic factor (BDNF). However, Ca2+ also activates calcineurin, inhibiting the BDNF-induced extracellular signal-regulated kinase (ERK) cascade. When the granule cells migrate to the internal granule layer during late postnatal stages, they are hyperpolarized and the blockade by calcineurin is removed, thereby enabling ERK-induced NR2C expression. On the other hand, how the gene expression of NR2B is down-regulated and eventually dominated by that of NR2A in other regions of the adult brain remains unknown (see > Section 2.3.4.).

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2.3 Trafficking Like other neurotransmitter receptors, NMDA receptors are mainly synthesized and assembled in the rough endoplasmic reticulum (ER), which is typically located in the soma of neurons. They are then transported to the Golgi apparatus and targeted to the dendrites via post-Golgi vesicular transport. The number and the composition of NMDA receptors are thought to be regulated by endocytotic and exocytotic processes.

2.3.1 ER Exit The C1 cassette of NR1 (> Figure 16-1) contains an ER retention signal (RRR); this signal must be masked by C-termini of NR2 subunits (Lau et al., 2007), NR3 subunits (Perez-Otano et al., 2001; Matsuda et al., 2003), or PDZ proteins (Standley et al., 2000) for the NMDA receptor complex to exit the ER. Conversely, NR2 and NR3 subunits are retained in the ER unless they form a complex with the NR1 subunit (PerezOtano et al., 2001; Matsuda et al., 2003). In addition, phosphorylation by protein kinase C (PKC) or protein kinase A near the ER retention signal of the NR1 subunit can release NMDA receptors from the ER (Scott et al., 2003). Similarly, a PDZ protein, SAP97 that binds to the C-terminus of NR2 is phosphorylated by CaMKIIa, allowing NMDA receptors to exit the ER (Mauceri et al., 2007). In contrast, a sustained increase in neuronal activity is shown to reduce the expression of the splice variant of NR1 containing the C20 cassette, which contains an ER export signal; this mechanism may serve as a homeostatic regulator to prevent the overstimulation of NMDA receptors (Scott et al., 2001).

2.3.2 Transport Along Dendrites While the polarized sorting of AMPA receptors to the dendritic domain is achieved by an adaptor complex, AP-4, NMDA receptors are properly sorted to the dendrites in mice lacking AP-4 (Matsuda et al., 2008). Thus, how NMDA receptors are sorted to the dendrites remains unclear. Once in the dendrites, NR2B is transported along microtubules by KIF17, a neuron-specific molecular motor, through an interaction with the PDZ proteins mLin-10 (Mint1), mLin-2 (CASK), and mLin-7 (MALS/Velis) (Setou et al., 2000). At extrasynaptic dendritic sites, CaMKII phosphorylates KIF17, causing the release of mLin-10 from KIF17 (Guillaud et al., 2008).

2.3.3 Exocytosis/Endocytosis at Synapses The exocyst complex is thought to direct intracellular membrane vesicles to fuse with the plasma membrane. SAP102, a PDZ protein that binds to the C-terminus of NR2, interacts with an exocyst protein, Sec8, thereby assisting the exocytosis of the NMDA receptor complex near synapses (Sans et al., 2003). Furthermore, PKC is known to phosphorylate SNAP25 and may also enhance the exocytosis of NMDAR-containing membrane vesicles (Perez-Otano et al., 2005). The juxtamembrane region of the cytoplasmic tail of NR1 contains sorting signals (YKRH and VWRK; > Figure 16-1) that target NMDA receptors to late endosomes and lysosomes for degradation (Scott et al., 2004). Similarly, the juxtamembrane regions of NR2 contain sorting signals (YWXX) for the degradation pathway. In contrast, the distal region of the C-terminus of NR2B contains a sorting motif (YEKL) that is recognized by an adaptor protein, AP-2, and targets NMDA receptors to recycling endosomes (> Figure 16-1) (Lavezzari et al., 2004). Similarly, AP-2 recognizes the di-leucine motif located in the C-terminus of NR2A, although this recognition is less effective than that for NR2B. Indeed, NMDA receptors containing NR2A are more stable than those containing NR2B. Interestingly, when the tyrosine residue in the YEKL motif of NR2B is phosphorylated by Fyn (Prybylowski et al., 2005) or when PSD-95 binds to a nearby region (Roche et al., 2001), the endocytosis of NMDA receptors is inhibited. However, in genetically engineered mice in which this motif was disrupted, the number of NR2B molecules on the cell

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surface was not significantly affected; instead, NR2B became localized in perisynaptic regions (Nakazawa et al., 2006). In addition, the overexpression of PSD-95 depresses the expression of NR2B at synapses (Losi et al., 2003). Thus, PSD-95 and tyrosine phosphorylation may have complex effects on NMDA receptor trafficking. The C-terminus of NR3A associates with PACSIN/sindapin1 (> Figure 16-1), which can recruit components of endocytic machinery, like dynamin-1 and clathrin, in an activity-dependent manner. Thus, NR3A may serve as an endocytic adaptor for NMDA receptors (Perez-Otano et al., 2006).

2.3.4 Synaptic Versus Extrasynaptic NMDA Receptors During development, the composition of synaptic NMDA receptors changes, switching from a predominance of NR2B-containing to NR2A-containing receptors. This switchover is bidirectionally regulated by neuronal activities, such as visual experience and odor discrimination (Quinlan et al., 2004). Since the changes in subunit composition occur in a matter of seconds and last for a least an hour (Bellone et al., 2007), the local endocytic and exocytic trafficking processes described above may underlie this phenomenon. In contrast, the developmental NR2B-NR2A switch proceeds much more slowly and may involve other mechanisms (Perez-Otano et al., 2005). Neuronal activities accelerate the turnover rate of several synaptic proteins, like NR2B and SAP-102, while keeping other components, like NR2A and PSD-95, constant (Ehlers, 2003). Such differential sensitivity to ubiquitin-based protein degradation may be one of the mechanisms responsible for the activity-dependent switchover of NMDA receptor subunits (PerezOtano et al., 2005). In contrast to the situation in immature neurons, the composition of NMDA receptors no longer changes after increased neuronal activities in mature neurons (Bellone et al., 2007). NR3A-containing NMDA receptors are mainly located at extrasynaptic sites, probably because synaptic activity induces the removal of NR3A-containing NMDA receptors through a PACSIN/syndapin1 pathway (Perez-Otano et al., 2006). However, NR3A-containing receptors are located at climbing fiber-interneuron synapses in the cerebellum (Fukaya et al., 2005b). In addition, NR3B-containing receptors, which probably do not bind to PACSIN/syndapin1, are also located at extrasynaptic sites (Matsuda et al., 2003). Thus, there may be additional mechanisms regulating the synaptic localization of NR3-containing NMDA receptors.

2.4 Functions The conventional mode of function of NMDA receptors is achieved through their Ca2+ permeable channels, which are blocked by Mg2+ in a voltage-dependent manner. When postsynaptic membranes are depolarized enough to relieve the blockade of NMDA receptors by Mg2+, presynaptically released glutamate can gate Ca2+ influx through the NMDA receptors, thereby enabling NMDA receptors to serve as a coincidence detector of presynaptic and postsynaptic activities (> Figure 16-2). Large increases in Ca2+ levels preferentially activate CaMKIIa, leading to the long-term potentiation (LTP) of AMPA receptor-mediated synaptic neurotransmission, whereas small increases activate calcineurin, followed by the long-term depression (LTD) of neurotransmission. For longer-term synaptic plasticity (> Figure 16-2), the expressions of several plastic genes, like BDNF, are orchestrated by the phosphorylation of cyclic AMP responsive elementbinding protein (CREB) and NF-kB (Rao et al., 2007).

2.4.1 Functional Diversities of NR1/NR2 Complexes NMDA receptors composed of different NR2 subunits exert distinct functions, probably reflecting differences in their desensitization kinetics (> Table 16-1), downstream signaling pathways, and subcellular localization. Some proteins, like CaMKIIa (Leonard et al., 1999), Rack1 (Yaka et al., 2002), and RasGRF1, a Ras-specific GDP/GTP exchange factor (Krapivinsky et al., 2003), preferentially bind to NR2B. In addition,

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although various PDZ proteins, like PSD-95 and SAP-102, bind similarly to NR2A and NR2B in vitro, PSD-95 and SAP-102 are preferentially found in the same complex as NR2A and NR2B, respectively, in vivo (Ehlers, 2003). Similarly, although the reason is unclear, NR2B-containing NMDA receptors are reported to associate preferentially with synaptic Ras GTPase activating protein (SynGAP) (Kim et al., 2005) and Rap1, a member of the Ras GTPase family (Zhu et al., 2005). In contrast, NR2A is preferentially associated with Ras and Rap2, a member of the Ras GTPase family, for unknown reasons. Although NR2Aand NR2B-containing NMDA receptors are differentially coupled to intracellular kinase activities like those of ERK, 38-kDa mitogen-activated protein kinase (p38 MAPK), and c-Jun NH2-terminal kinase (JNK), their roles in modulating LTP and LTD are not completely clear. For example, at the CA1 region of the hippocampus, LTP was reported to be induced specifically by NR2A-containing NMDA receptors (Liu et al., 2004) and NR2B-containing NMDA receptors (Barria et al., 2005) or nonselectively by both NR2Aand NR2B-containing receptors (Massey et al., 2004; Berberich et al., 2007). Similarly, the exact subunit composition of synaptic versus extrasynaptic NMDA receptors is still controversial. For example, NR2Aand NR2B-containing NMDA receptors are reported to be located differentially to synaptic and extrasynaptic sites, respectively (Barria et al., 2002; Nagy et al., 2004), but these receptors have also been found at both sites nonselectively (Thomas et al., 2006b).

2.4.2 Functions of NR3-Containing NMDA Receptors As described in > Section 2.1., NR3 subunits reduce the amplitudes and Ca2+ permeability of glutamateinduced currents in NMDA receptors (Perez-Otano et al., 2001; Matsuda et al., 2003). After brain injury, NR3A is expressed in oligodendrocyte processes and may form NMDA receptors with low Ca2+permeability (Salter et al., 2005). Similarly, mature motoneurons, which express NMDA receptors containing NR3B subunits, exhibit very small NMDA-induced currents (Hori et al., 2002). NMDA receptors composed of only NR1 and NR3A or NR1 and NR3B subunits exhibit unique excitatory glycine responses in Xenopus oocytes; the glycine response was evoked by glycine alone and inhibited by D-serine, but not by AP-5 or picrotoxin (Chatterton et al., 2002). However, NMDA receptors composed of NR1 and NR3 subunits do not exhibit the glycine response in mammalian heterologous cells (Nishi et al., 2001; Matsuda et al., 2003; Smothers et al., 2007). Interestingly, when equal amounts of NR1, NR3A, and NR3B cDNAs were cotransfected, the glycine response was observed in mammalian cells (Smothers et al., 2007). Although the NR2 to NR3B switchover of NMDA receptors is observed in early postnatal motoneurons, NR3A is expressed at very low levels in these neurons (Fukaya et al., 2005a). Thus, excitatory glycine receptors with similar amounts of NR1, NR3A, and NR3B are unlikely to occur in motoneurons. Therefore, whether NR3 forms an excitatory glycine receptor in vivo remains unclear.

3

Delta Receptors

3.1 Structure and Basic Properties The d family of iGluR, consisting of GluRd1 and GluRd2, is equidistant from the NMDA receptors and the AMPA receptors. Thus, the membrane topology of GluRd1 and GluRd2 is thought to be similar to that of other iGluRs (> Figure 16-3). Similarly, GluRd1 and GluRd2 are thought to form a tetramer. However, the vast majority of GluRd2 and, possibly, GluRd1 are thought to exist as a homomeric receptor in vivo (Yuzaki, 2008). Recently, the crystal structure of the LBD of GluRd2 has been solved. D-serine and glycine bind to the purified LBD of GluRd2 with low affinities and KD values of 1.1 and 3.6 mM, respectively (Naur et al., 2007). Like the LBD of other iGluRs, the apo structure of GluRd2 shows a twofold symmetric dimer in which each monomer forms a typical bi-lobed structure (> Figure 16-3). A unique feature of the GluRd2 LBD is the tight binding of two Ca2+ ions at the dimer interface. Ligand binding induces a domain closure

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. Figure 16-3 Presumed membrane topology and associated intracellular molecules of GluRd2. The ligand-binding domain (LBD) binds to D-serine and glycine with the indicated affinities. Ca2+ ions bind to the dimer interface and may modulate the gating process. The ion selectivity filter region in the TM2 is encoded by glutamine (Q). Intracellular molecules that bind to the C terminus of GluRd2 and their putative functions are shown. PDZ proteins bind to the TSI motif at the end of the C-terminus (Adapted from Yuzaki, 2008)

of about 30 , which is much larger than that observed in the LBD of AMPA receptor GluR2 (21 ). Similarly, Cl ions bind to the dimer interface and modulate the desensitization kinetics of kainate receptors (Plested et al., 2007).

3.2 Genes and Expression The size of the genes encoding GluRd1 and GluRd2 are much larger than those of other iGluRs in both mice and humans. Indeed, the gene encoding GluRd2 is the 13th largest known human gene. Neither GluRd1 nor GluRd2 undergoes any known process of alternative splicing or RNA editing. During the early postnatal period, GluRd1 mRNA is transiently expressed at higher levels in the caudate and thalamic nuclei. It is continuously expressed at low levels in the pyramidal and dentate granule cell layers of the hippocampus during development and in adulthood. GluRd1 is also highly expressed in the inner hair cells in adult rats and the ganglion and bipolar cells of the retina (Jakobs et al., 2007). GluRd2 is highly and predominantly specifically expressed in a major cell type of the cerebellum, the Purkinje cell. GluRd2 mRNA can be detected in Purkinje cells in mice as early as embryonic day 15, increases markedly during the 2nd and 3rd weeks of postnatal development, and remains high throughout adulthood. However, it is also expressed at low levels in several neurons in the midbrain-spinal cord region, like the dorsal cochlear nucleus and the trigeminal motor nucleus, as well as in the pineal gland.

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3.3 Trafficking Unlike other iGluRs, which are often found in the ER and extrasynaptic regions, GluRd2 proteins are efficiently transported to the cell surface and targeted to the spines along the distal dendrites of cerebellar Purkinje cells, where parallel fiber (PF) axons from granule cells form synapses. Similarly, unlike AMPA receptors, which are dynamically endocytosed or exocytosed during LTD or LTP, respectively, GluRd2 is relatively stable at the PF synapses. The GluRd1 trafficking mechanism is unknown.

3.3.1 ER Exit and Transport along Dendrites The efficient cell-surface transport of GluRd2 requires its C-terminal juxtamembrane region of 13 amino acids (Matsuda et al., 2004), to which unknown factors may bind (> Figure 16-3). Various small in-frame deletions in the NTD of GluRd2 found in various hotfoot mutant mice impair the assembly of GluRd2 and its subsequent exit from the ER (Matsuda et al., 2002; Wang et al., 2003). These findings indicate that the NTD of GluRd2 is essential for receptor assembly and that unstable oligomers may be retained in the ER by a quality control mechanism. GluRd2 is sorted to the dendrites by AP-4 (Matsuda et al., 2008), which directly binds to the middle region of the GluRd2 C-terminus (> Figure 16-3).

3.3.2 Exocytosis/Endocytosis at Synapses The stable expression of GluRd2 at synapses requires the middle region of the 12 C-terminal amino acids (E-region; > Figure 16-3) (Matsuda et al., 2006b). Conversely, the addition of the E-region to the C-terminus of GluR1 AMPA receptors significantly increased the amount of GluR1 at synapses. Similarly, the amount of GluRd2 proteins at PF synapses was significantly decreased in mice that expressed GluRd2 lacking the E-region (Yasumura et al., 2008). Although a PDZ protein Shank binds to the region near the E-region (Uemura et al., 2004), the factor responsible for the stabilization of GluRd2 at synapses remains unclear. The interaction of GluRd2 with spectrin may also be required for its stable expression. GluRd2 is not normally expressed at spines located on the proximal dendrites of Purkinje cells, where climbing fibers form synapses. When the electrical activity of the climbing fibers is blocked, GluRd2 can be observed in the proximal dendrites. It is hypothesized that climbing fiber-induced Ca2+ spikes may increase the intracellular Ca2+ concentration and destabilize the spectrin–GluRd2 association at proximal dendrites (Hirai, 2001). Similarly, a mutation of b-III spectrin, which is highly expressed in Purkinje cells, is reported to cause the loss of GluRd2 at synapses in families with spinocerebellar ataxia type 5 (Ikeda et al., 2006).

3.4 Functions Mice lacking GluRd1 show grossly normal innervation patterns of the inner hair cells and functions of the hippocampus and vestibular system. However, the cochlear threshold for frequencies higher than 16 kHz is shifted significantly and mice are more susceptible to acoustic injury (Gao et al., 2007). These findings suggest a modulatory role of GluRd1 in high-frequency hearing, but the underlying mechanisms remain unclear. Detailed morphological analyses of GluRd2-null mice (i.e., hotfoot and genetically engineered GluRd2 knockout mice) have revealed two major functions of GluRd2 at PF–Purkinje cell synapses (> Figure 16-4): the formation/maintenance of PF–Purkinje cell synapses and the regulation of LTD (Yuzaki, 2004).

3.4.1 Phenotypes of Glurd2-Null Mice The number of PF–Purkinje cell synapses is markedly reduced in GluRd2-null cerebella. Approximately 40% of the spines remain free of any presynaptic contact in GluRd2-null cerebella. By contrast, free spines,

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. Figure 16-4 Major cerebellar phenotypes of GluRd2- and Cbln1-null mice. GluRd2- and Cbln1-null mice show ataxic gait, motor discoordination, and defective motor learning. Cbln1 is produced and released from granule cells, while GluRd2 is expressed at the dendritic spines of Purkinje cells. GluRd2- and Cbln1-null mice show two specific and characteristic electron microscopic anomalies at parallel fiber–Purkinje cell synapses: numerous free spines not innervated by parallel fibers, and a postsynaptic density (PSD) that is disproportionally longer than that of the opposing presynaptic active zone length. In addition, a stimulus that normally evokes the long-term depression (LTD) of parallel fiber transmission failed to induce the endocytosis of AMPA receptors (Adapted from Yuzaki, 2008)

which are observed when granule cells are damaged by irradiation or other genomic mutations, are transient and eventually innervated by the remaining PFs. In addition, the remaining PF–Purkinje cell synapses in GluRd2-null mice frequently exhibit a postsynaptic density (PSD) that is disproportionally longer than that of the opposing presynaptic active zone (> Figure 16-4). These findings indicate that GluRd2 plays a unique role in aligning and maintaining the PSD with the presynaptic element at PF– Purkinje cell synapses. Increased conjunctive activities of PFs and CFs induce LTD of PF responses in wild-type Purkinje cells; LTD is thought to underlie some forms of motor learning. LTD is completely abrogated in GluRd2-null Purkinje cells. Conversely, the application of an antibody against the LBD of GluRd2 caused the endocytosis of synaptic AMPA receptors and the LTD of PF–Purkinje cell transmission (Hirai et al., 2003). These findings indicate that GluRd2 actively controls AMPA receptor endocytosis, thereby modifying the synaptic plasticity at PF-Purkinje cell synapses. Interestingly, all the behavioral, physiological, and anatomical phenotypes of GluRd2-null mice are shared by Cbln1-null mice (> Figure 16-4). In addition, mice that lacked both GluRd2 and Cbln1 did not show an additive phenotype, but rather were similar to mice lacking only GluRd2. These findings suggest that Cbln1, which is expressed and released from granule cells, and GluRd2 engage in a common signaling pathway or process crucial for synapse formation/maintenance and plasticity.

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3.4.2 Mechanisms Clarified by the “Transgenic Rescue” Approach The mechanism by which GluRd2 achieves the above-mentioned functions has been unclear, mainly because of the lack of specific pharmacologic tools. To obtain clues to the functioning of GluRd2 without relying on pharmacological tools, we have recently exploited the “transgenic rescue” approach (Yuzaki, 2005). A series of mutant GluRd2 transgenes were introduced into GluRd2-null Purkinje cells using either transgenic mice or virus vectors. If the mutant GluRd2 transgene was as effective as the wild-type GluRd2 transgene in rescuing the phenotypes of the GluRd2-null mice described in > Section 3.4.1., the mutated regions were judged as functionally unimportant (> Table 16-2). For example, because a mutant GluRd2 transgene (d2-R/K), in which the glutamate-binding motif conserved in iGluRs was mutated, rescued all the abnormal phenotypes of GluRd2-null mice, the binding of glutamate analogs is not essential for these functions of GluRd2 in vivo (Hirai et al., 2005). Because D-serine does not bind to this mutant GluRd2, the functional significance of the binding of D-serine remains to be elucidated. It has been a long-standing question whether GluRd2 serves as an ion channel, like other iGluRs. Support for GluRd2 channel activity has come from studies on spontaneously occurring ataxic mutant lurcher mice. A point mutation at the end of TM3 of GluRd2 causes the continuous activation of its channels (GluRd2Lc). Like AMPA and kainate receptors, GluRd2Lc exhibited a rectified current-voltage relationship and moderate Ca2+ permeability (Kohda et al., 2000; Wollmuth et al., 2000). In addition, similar to wild-type AMPA receptors, the Ca2+-permeability of GluRd2Lc was abolished by the substitution of glutamine with arginine at the putative channel pore region (> Figure 16-3). These findings suggest that GluRd2Lc forms an ion channel with distinct properties. However, a mutant GluRd2 transgene (d2-Q/R), in which the ion selectively filter conserved in iGluRs was mutated, successfully rescued all the abnormal phenotypes of GluRd2-null mice. (Kakegawa et al., 2007a) (> Table 16-2). Furthermore, a mutant GluRd2 transgene (d2-V/R), in which valine located one position upstream of the Q/R site was replaced with arginine, also completely rescued LTD (Kakegawa et al., 2007b) (> Table 16-2). Since similar mutations disrupt the channel pores of AMPA and kainate receptors, as well as those formed by GluRd2Lc, these findings indicate that...” these findings indicate that although GluRd2 belongs to the iGluR family, it does not function as a channel, at least with regard to the regulation of LTD and PF synapse formation/ maintenance. In this case, although GluRd2Lc forms an ion channel, the ability of wild-type GluRd2 to gate current may simply be a function that was lost during evolution. Interestingly, a mutant GluRd2 transgene lacking the C-terminal 7 amino acids, completely disrupting the PDZ-binding motif (d2-DCT7), could not restore abrogated LTD at PF–Purkinje cell synapses and motor

. Table 16-2 Rescue of phenotypes of GluRd2-null mice by mutant GluRd2 transgenes (Adapted from Yuzaki, 2008)

Genotype

GluRd2null mice a

GluRd2 TG introduced into Purkinje cells none +d2-WT +d2-R/K +d2-Q/R +d2-V/R +d2-DCT7

Parallel fiber synapses ↓↓↓a ! ! ↓c n.e. !

Climbing fiber synapses ↑↑↑ ! ! ! n.e. !

Long-term depression ↓↓↓ ! ! ! ! ↓↓d

Gait and rotarod performance ↓↓↓ ! ! ! n.e. ↓e

Motor learning ↓↓↓ ! n.e.b n.e. n.e. ↓↓↓

Downward arrows indicate reduced parallel fiber synapses, impaired long-term depression, dyscoordinated gait/rotarod, or impaired motor leaning; rightward arrows indicate normal phenotypes; upward arrows indicate sustained innervation of Purkinje cells by supernumerous climbing fibers. b n.e.: not examined c Not completely rescued; however, the expression level of d2-Q/R was not as high as that of d2-WT d Only partially rescued e Performances on rotorod tests at 10 r.p.m, but not at 20 r.p.m., were rescued

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. Figure 16-5 Working hypothesis for the function of GluRd2. The control of AMPA receptor trafficking and LTD does not require ion channel activities but does depend on nonionotropic functions via C-terminal associated proteins. The N-terminal domain of GluRd2 may bind directly or indirectly to some adhesion molecules located at presynaptic sites to regulate the morphological integrity at parallel fiber synapses. The function achieved by binding to D-serine is currently unclear (Adapted from Yuzaki, 2008)

learning (Kohda et al., 2007; Uemura et al., 2007; Kakegawa et al., 2008). Thus, molecules that bind to the C-terminal 7 amino acids of GluRd2 play a crucial role in LTD induction and motor learning (> Figure 16-3). For example, delphilin is reported to bind to Src tyrosine kinase via the FH1 domain (Miyagi et al., 2002; Matsuda et al., 2006a), and PTPMEG contains a protein tyrosine phosphatase domain. Because cerebellar LTD depends on the tyrosine phosphorylation status in Purkinje cells, GluRd2’s C-terminus may regulate LTD induction through an interaction with delphilin and PTPMEG (Kina et al., 2007). In contrast, d2-DCT7 successfully rescued the morphological integrity at PF synapses (> Figure 16-4), such as free spines at PF synapses (Kakegawa et al., 2008). Thus, these functions may be differentially regulated by other regions of GluRd2 (> Figure 16-5). Recently, the N-terminal extracellular domain of AMPA receptors has been shown to bind to an adhesion molecule, N-cadherin (Nuriya et al., 2006; Saglietti et al., 2007). Therefore, the N-terminal domain of GluRd2 may also bind directly or indirectly to some adhesion molecules located at presynaptic sites (> Figure 16-5).

4

Conclusion

AMPA receptors are “executers” of fast excitatory neurotransmission, whereas NMDA receptors and GluRd2 can be regarded as “regulators” that control AMPA receptor trafficking and synaptic plasticity. Intuitively, GluRd2 is predominantly expressed at the PF–Purkinje cell synapse, which is one of the very rare regions where no functional NMDA receptors are expressed. Thus, at PF synapses, GluRd2 might exert some of the functions that are normally achieved by NMDA receptors in other brain regions. For example, D-serine has been recently shown to serve as a “gliotransmitter,” which is released from glia and activates neuronal NMDA receptors to regulate LTP or LTD in hippocampal neurons (Martineau et al., 2006; Billard, 2008). Although D-serine binding to GluRd2 was not required for the major functions of GluRd2, it might serve to regulate metaplasticity under certain conditions. It has become increasingly clear that signaling

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mechanisms via intracellular proteins that bind to the C-termini of GluRd2 and NMDA receptors play key roles in regulating the trafficking of AMPA receptors. Therefore, characterization of the molecular mechanisms activated by GluRd2 and NMDA receptors should add to our knowledge of each receptor family, facilitating a better understanding of the synaptic plasticity underlying learning and memory.

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Shank-binding segment of glutamate receptor delta2 in synaptic localization and cerebellar functions. Neurosci Lett 433: 146-151. Yuzaki M. 2003. The delta2 glutamate receptor: 10 years later. Neurosci Res 46: 11-22. Yuzaki M. 2004. The delta2 glutamate receptor: A key molecule controlling synaptic plasticity and structure in Purkinje cells. Cerebellum 3: 89-93. Yuzaki M. 2005. Transgenic rescue for characterizing orphan receptors: A review of delta2 glutamate receptor. Transgenic Res 14: 117-121. Yuzaki M. 2008. New (but old) molecules regulating synapse integrity and plasticity. Cbn1 and the d2 glutamate receptor. Neurosci, in press. Zhu Y, Pak D, Qin Y, McCormack SG, Kim MJ, et al. 2005. Rap2-JNK removes synaptic AMPA receptors during depotentiation. Neuron 46: 905-916.

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Y. Kubo . M. Tateyama

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334

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Sensitivity to Polyvalent Cations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334

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Crystal Structure Analysis—Dimeric Structure and the Venus Flytrap Module . . . . . . . . . . . . . . . 335

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Crystal Structure Analysis—Binding Sites for Polyvalent Cations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 336

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Structural Rearrangement of the Cytoplasmic Region of mGluR1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 336

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Factors Regulating the Switching Among Multiple Signaling Pathways 1—Ligand Type . . . . . . 338

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Factors Regulating the Switching Among Multiple Signaling Pathways 2—Differences Between Splice Variants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339

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Factors Regulating the Switching Among Multiple Signaling Pathways 3—Effects of Cytoplasmic-Binding Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 340

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Physiological Significance of Gd3+ Sensing Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341

10 Future Directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 342

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Abstract: In this chapter, we focus chiefly on our studies of the regulatory mechanisms and structural rearrangements that affect signaling via the metabotropic glutamate receptor (mGluR), and we also introduce related studies by other groups investigating the dynamic aspects of mGluR1. The topics covered include (1) the sensitivity of mGluR1 to polyvalent cations, (2) fluorescence resonance energy transfer analysis of the structural rearrangement of the cytoplasmic region of mGluR1, and (3) switching of mGluR1 signaling among multiple pathways, depending upon the ligand type, the splice variation of the C-terminal structure, and the presence of cytoplasmic-binding proteins. List of Abbreviations: Cao2+, extracellular calcium; Ca2+ i , intracellular calcium; CFP, cyan fluorescent protein; FRET, fluorescence resonance energy transfer; Gdo3+, extracellular gadorinium; LTD, long-term depression; LTP, long-term potentiation; mGluR, metabotropic glutamate receptor; 7TMD, seven transmembrane domain; TIRF, total internal reflection; VFTM, venus flytrap module; YFP, yellow fluorescent protein

1 Introduction The existence of G protein-coupled, or metabotropic, glutamate receptors was first advocated based on functional evidence (Sugiyama et al., 1987). Within several years thereafter, the first cDNA encoding mGluR1 was isolated (Houamed et al., 1991; Masu et al., 1991), after which additional cDNA clones from the same family were isolated and classified into three subgroups: group 1 includes mGluR1 and 5, group 2 includes mGluR2 and 3, and group 3 includes mGluR4, 6, and 7. Subsequent functional analyses showed that group 1 receptors are coupled to Gq, while receptors in groups 2 and 3 are coupled to Gi/o (Nakanishi et al., 1994). Histochemical analyses showed these receptors to be expressed in distinct patterns in several areas of the brain, including in the hippocampus, cerebellum, and olfactory bulb. The physiological significance of mGluR1 was demonstrated in gene knockout studies that showed that plastic changes in the efficiency of synaptic transmission, such as long-term potentiation (LTP) and long-term depression (LTD), were lost in mGluR1 knockout mice (Aiba et al., 1994a, b; Kano et al., 1995). For that reason, mGluR1 is thought to be a key molecule involved in memory and learning. mGluR1 is classified as a family-C G protein-coupled receptor, along with receptors for GABA (Kaupmann et al., 1997, 1998), Ca2+, sweet and umami tasting substances, and various pheromones (reviewed in Mitri et al., 2004). The characteristic structural feature of mGluR1 and other family-C receptors is their large extracellular N-terminal region, and a crucial breakthrough in this area of research was resolution of the X-ray crystal structure of part of the extracellular domain of mGluR1, which showed the receptor to be a dimer (Kunishima et al., 2000; Tsuchiya et al., 2002). This finding and other related works on the structure and function of homo- and heterodimeric G protein-coupled receptors have been reviewed extensively (De Blasi et al., 2001; Hermans and Challiss, 2001; Jensen et al., 2002; Jingami et al., 2003; Pin et al., 2003, 2004, 2005; Kubo and Tateyama, 2005). More recently, the structures of the extracellular regions of mGluR3 and mGluR7 were also solved and provide a structural basis for the subtype-specific binding of agonists (Muto et al., 2007). We joined this research when we found that mGluR1 is sensitive to extracellular polyvalent cations (Kubokawa et al., 1996; Kubo et al., 1998). After that, we investigated the dynamic structural rearrangement of the cytoplasmic region upon activation by fluorescence resonance energy transfer (FRET) analysis under the total internal reflection (TIRF) microscope (Tateyama et al., 2004). We also studied the mechanisms involved in regulating mGluR1-mediated G protein signaling, which is dependent upon the ligand type (Tateyama and Kubo, 2006), the splice variant involved, and the presence of cytoplasmic-binding proteins (Tateyama and Kubo, 2007). In this chapter, we will focus chiefly on our studies of mGluR1, and we will also introduce related studies by other groups investigating the dynamic aspects of mGluR1.

2 Sensitivity to Polyvalent Cations While attempting to isolate cDNA encoding a certain peptide hormone receptor from salmon brain using a Xenopus oocyte expression system, we happened to notice that there was a cDNA library pool that induced

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basal Gq activity (i.e., a sustained Ca2+–Cl
current), even before application of the hormone. By repeatedly subdividing that cDNA library, we were ultimately able to isolate a single cDNA clone. Upon determining its sequence, the isolated cDNA was unexpectedly identified as a salmon orthologue of mGluR1 (Kubokawa et al., 1996). Moreover, because it induced a marked sustained Gq activity in the absence of glutamate, we speculated that a substance in the bath solution was stimulating it. At that time, a cDNA clone encoding the extracellular Ca2+ receptor (CaR) was isolated and was shown to belong to the same family-C as mGluR1 and to be characterized by a long extracellular N-terminal region (Brown et al., 1993). Given that CaR and mGluR1 belong to the same family and that CaR can be activated by extracellular Ca2+(Cao2+), we speculated that it might be the Ca2+ in the bath solution that induced the persistent activation of salmon mGluR1. Consistent with that idea, the basal activity subsided when Cao2+ was removed from the extracellular solution and was triggered by reintroducing Cao2+. We later observed that rat mGluR1 is also sensitive to Cao2+ and, in a mutagenesis study, showed that Ser166, located close to the glutamate-binding site, is critical for Cao2+ sensitivity. We therefore concluded that mGluR1 can be activated by Cao2+ in Xenopus oocytes (Kubo et al., 1998). Moreover, similar to CaR (Brown et al., 1993), mGluR1 in Xenopus oocytes proved highly sensitive to the trivalent cations Gd3+, Tb3+, and La3+ (Kubo et al., 1998). Based on analysis of inositol phosphate turnover in CHO or BHK cells, it was suggested that Cao2+ does not act directly as an mGluR1 agonist (Saunders et al., 1998; Nash et al., 2001). On the other hand, using FRET analysis with HEK 293 cells, we observed that exposure to 5-mM Ca2+ induced structural rearrangement of mGluR1, and that the FRET change induced by Cao2+ was abolished by an S166D mutation (Tateyama et al., 2004), which suggests that Cao2+ does indeed act as an agonist. But as the sensitivity of mGluR1 to Cao2+ in cultured mammalian cells is substantially lower than that previously observed in oocytes, the physiological role of Cao2+ as an mGluR1 agonist may be minor, as compared with its role as a CaR agonist. The major physiological role of Cao2+ via mGluR1 is more likely modulation of glutamateinduced responses (Saunders et al., 1998; Nash et al., 2001). Cao2+ reportedly enhances (Saunders et al., 1998) and sustains (Miyashita and Kubo, 2000a; Nash et al., 2001) glutamate response mediated by mGluR1. In primary cultured cerebellar Purkinje cells, Cao2+ modulates the dynamic range of the glutamate dose-response via mGluR1—that is, Cao2+ increases the sensitivity to low concentrations of glutamate (Tabata et al., 2002).

3 Crystal Structure Analysis—Dimeric Structure and the Venus Flytrap Module The structure of part of the extracellular domain of mGluR1 has been solved by X-ray crystallography (Kunishima et al., 2000; Tsuchiya et al., 2002). Those studies clearly showed that mGluR1 is a homodimeric molecule; that each subunit has an upper and a lower lobe, later named the venus flytrap module (VFTM); and that the glutamate-binding site is located in the center of the two lobes. Two dynamic structural rearrangements have been shown to occur upon glutamate binding: closure of the VFTM in each subunit and movement of the two subunits toward one another. The first change is called the ‘‘Open’’ to ‘‘Closed’’ transition of the VFTM, while the second is called the ‘‘Resting’’ to ‘‘Activated’’ transition (Kunishima et al., 2000). The ‘‘Open–Open’’ to ‘‘Closed–Open’’ transition, a closure of one of the two VFTMs, reportedly can occur even in the absence of glutamate, though the binding of glutamate facilitates a shift in the equilibrium from the ‘‘Open–Open’’ to ‘‘Closed–Open’’ configuration (Kunishima et al., 2000). It is noteworthy that a crystal structure in the ‘‘Closed–Closed’’ configuration has never been obtained, even in the presence of glutamate, and it has been suggested that the two subunits do not close simultaneously, presumably due to an intersubunit-negative allosteric interaction. In that regard, Suzuki et al. (2004) measured changes in intrinsic tryptophan fluorescence to test whether there is an allosteric interaction between the dimeric ligand-binding sites on mGluR1, and were able to estimate the concentration–binding relationship over a wide range of glutamate concentrations. The Hill coefficient of the saturation-binding curve indicated a strong negative cooperativity affecting glutamate binding to the dimer’s two binding domains. Based on that finding, they suggested that the ‘‘Closed–Open’’ conformation with one glutamate bound may be the standard active state. On the other hand, Kniazeff et al. (2004) used

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various chimeric molecules, in which the mGluR1 extracellular domain was incorporated into the seven transmembrane domain (7TMD) and cytoplasmic domain of GABAB1R or GABAB2R, to show that the ‘‘Closed–Open’’ state is only a partial activation, and that the ‘‘Closed–Closed’’ state is necessary for full activity. Similarly, by characterizing the properties of the wild-type homodimer and a Y74A mutant, which was much less sensitive to glutamate (Kammermeier and Yun, 2005), showed that glutamate binding to only one subunit of the dimer is not sufficient to activate mGluR1. The difference in the findings of these studies might in part reflect the differences in the experimental approaches: a functional analysis of the whole protein vs. a biochemical analysis using recombinant protein fragments. Alternatively, it may be that an endogenous molecule that enables dual closure is present in the expression systems used, but is not included with the purified recombinant protein.

4 Crystal Structure Analysis—Binding Sites for Polyvalent Cations Influenced by our study showing polyvalent cation sensitivity of mGluR1, Jingami’s group tried to identify the cation-binding site in the crystal with the aim of better understanding the polyvalent cation sensitivity of mGluR1 from a structural point of view. Notably, when crystals were grown in the presence of Ca2+, a bound Ca2+ was not observed at Ser166, its presumed binding site (Kunishima et al., 2000). This might reflect the low affinity of Ca2+ (EC50 ffi 5 mM) for its binding site and a binding mode that differs from the high-binding affinity of Ca2+—for example, to calmodulin. By contrast, the binding of the trivalent cation Gd3+ was readily confirmed in crystal. The binding site was different from that of either Ca2+ or glutamate, and located at an intersubunit slit between the two lower lobes, where negatively charged amino acids are clustered (Tsuchiya et al., 2002). We further confirmed the interaction of Gd3+ with this region in a mutagenesis study. By introducing an E238Q mutation in the center of the binding site, the response to Gd3+ was completely eliminated without changing the sensitivity to glutamate or Ca2+ (Abe et al., 2003b). It is noteworthy that in the presence of glutamate and Gd3+, the structure of mGluR1 assumes the Closed–Closed/Activated configuration (Tsuchiya et al., 2002). This suggests that the negative allosteric interaction inhibiting dual closure of the VFTMs upon glutamate binding is caused by the repulsion of clusters of negatively charged amino acids at the dimer interface and is mitigated by Gd3+. The binding of Gd3+ within the cluster apparently neutralizes to some degree the negative charge, thereby reducing the repulsion between the subunits enough to enable dual closure of the VFTMs. Consistent with that idea is the finding that neutralization of E238 by substituting it with His or Ala leads to an increase in basal mGluR1-mediated activity (Jensen et al., 2001, 2002). This may also explain why the Closed–Closed/ Activated configuration has not been structurally identified in the absence of Gdo3+, despite the high-affinity binding of glutamate.

5 Structural Rearrangement of the Cytoplasmic Region of mGluR1 The extracellular region beyond the VFTM and the transmembrane and cytoplasmic domains of mGluR1 were not included in the previously solved crystal structure. Bearing in mind that conformational changes in the cytoplasmic region are important for the function of G protein-coupled receptors, we used FRET analysis to examine the conformational changes in the cytoplasmic region of mGluR1 (Tateyama et al., 2004) (> Figure 17‐1). We initially coexpressed fusion proteins comprised of mGluR1 subunit linked to Cyan or Yellow Fluorescence Protein (CFP and YFP, respectively) in HEK293 cells. Because mGluR1 is dimeric, 50% of the expressed receptors were expected to have both CFP and YFP within the dimer. To efficiently detect the conformational changes upon glutamate binding, it was necessary to record only the fluorescent signals of mGluR1 expressed on the cell membrane. When we carried out FRET analysis under a TIRF microscope, we observed that upon application of glutamate, intersubunit FRET between the second short internal loops increased, whereas that between the first internal loops decreased. We also analyzed intrasubunit FRET and found that the relative positions of the truncated C-terminal cytoplasmic tail and

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. Figure 17‐1 FRET analysis of the structural rearrangement of the cytoplasmic domain of mGluR1 elicited by glutamate application. (a) The location of CFP and YFP in each construct, and respective data showing the changes in FRET values occurring upon glutamate application. (b) Postulated conformational change in the cytoplasmic domain of mGluR1. Circles show the 7TMD based on the structure of Rhodopsin. The relative position of the two subunits changes as a result of sliding and/or rotation, so that the distance between loops 1 (light gray) increases (a, left), while that between loops 2 (dark gray) decreases (a, right). No clear intrasubunit change in structure was detected (a, right). (Modified from original figures in Tateyama et al. (2004), with permission from Nature.)

the second short internal loop did not change significantly upon glutamate application (Tateyama et al., 2004). Thus, the change in the relative position of the two subunits was more pronounced than the conformational change within each subunit. Moreover, it was not simply that two subunits moved closer together, as the distance between first loops actually increased, while that between the second loops decreased. This suggests that a sort of rotation or shifting movement occurred (> Figure 17‐1). That a conformational change within each subunit could not be detected was unexpected, and further analysis seems warranted; for example, additional constructs could be used to obtain sufficient information to draw conclusions about intrasubunit movement (Parnot and Kobilka, 2004). The mechanisms linking

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agonist binding to conformational changes in the transmembrane and cytoplasmic regions are also of interest. Using evolutionary trace analysis and mutagenesis, it was recently shown that a conserved interdomain disulfide bridge is required for this allosteric interaction (Rondard et al., 2006; Muto et al., 2007).

6 Factors Regulating the Switching Among Multiple Signaling Pathways 1—Ligand Type It is generally accepted that mGluR1 links to the activation of Gq protein, which mediates activation of protein kinase C and increases in [Ca2+]i. However, there are also reports showing that mGluR1 links to the activation of Gs protein (Aramori and Nakanishi, 1992; Joly et al., 1995; Miyashita and Kubo, 2000b; Selkirk et al., 2001). As described in the previous section, different ligands (e.g., glutamate and Gd3+) bind to different regions of mGluR1, perhaps causing different activation conformations. We hypothesized that the G protein coupling and the downstream signaling might also vary depending on ligand type. To test this idea, we made simultaneous optical recordings of Gq- and Gs-mediated responses following activation of mGluR1a expressed in CHO cells (> Figure 17‐2). We monitored the Gs-coupled signaling (i.e., an increase . Figure 17-2 Gd3+ binding to mGluR1a activates signaling via Gq but not Gs. (a) Functional coupling of mGluR1a to Gq (changes in [Ca2+]i, abscissa) and Gs (normalized [cAMP]i, ordinate) signaling induced by 1-mM glutamate (open circles) or 100-mM Gd3+ (filled circles). (b, c) Glutamate dose–response relations for Gq (b) and Gs (c) signaling are shown as changes in [Ca2+]i (b) and normalized [cAMP]i (c), respectively. (Modified from original figures in Tateyama and Kubo (2006), with permission from Proc. Natl. Acad. Sci. USA.)

in [cAMP]i) using a FRET-based [cAMP] sensing protein developed by Zaccolo et al. (2000). To avoid overlap of the excitation wavelengths, we used Indo-1 to monitor the Gq-coupled signaling (i.e., increases in [Ca2+]i). As a control, we first examined the responses of the m1 muscarinic ACh receptor and the A2a adenosine receptor, which are known to be coupled exclusively to Gq and Gs, respectively. In these cases, only the expected Gq or Gs responses were observed, which confirmed the validity of the recording system. When glutamate was applied to CHO cells expressing mGluR1a, increases in both [Ca2+]i and [cAMP]i were observed. At the single cell level, there were cells that predominantly showed Gq or Gs responses, which might reflect cell-to-cell variation in the cellular conditions (e.g., the phosphorylation or redox state). On the whole, however, it was apparent that mGluR1a activation by glutamate is coupled to both Gq and Gs pathways. By contrast, when mGluR1a was activated by Gd3+, only Gq-coupled responses were observed; no Gs responses were detected (> Figure 17‐2). Taken together, these findings indicate that the specific nature of downstream mGluR1a signaling is ligand dependent (Tateyama and Kubo, 2006). Receptors are generally thought of as simple on–off switches, but this is not the case with mGluR1,

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which appears to be a switchable regulator of multiple signaling pathways (> Figure 17‐3). Similar observations of allosteric modulators and multiple conformations and signaling pathways also have been reported for other G protein-coupled receptors (Hermans and Challiss, 2001; Baker and Hill, 2007; Kobilka and Deupi, 2007; Leach et al., 2007). . Figure 17‐3 Activated states of mGluR1a Upon binding glutamate, mGluR1 goes from an Open–Open/Resting state to a Closed–Open/Activated state. In the presence of glutamate and Gd3+, and probably also Gd3+ alone, mGluR1 is able to go to a second activated state, the Closed–Closed/Activated state, because Gd3+ neutralizes negative charges clustered at the dimer interface, thereby mitigating their negative allosteric effect (Kunishima et al., 2000; Tsuchiya et al., 2002)

7 Factors Regulating the Switching Among Multiple Signaling Pathways 2—Differences Between Splice Variants Signaling by mGluR1a and mGluR1b, a splice variant with a shortened C-terminal cytoplasmic region, reportedly differs (Joly et al., 1995). To examine this difference in detail, we made simultaneous recordings of [Ca2+]i and [cAMP]i levels and compared the glutamate responses mediated via mGluR1a and 1b expressed in CHO cells (> Figure 17‐4a, b). We found that application of glutamate to cells expressing mGluR1b evoked Gq responses, with little or no Gs response. When we then compared the properties of several deletion mutants to identify the structural determinant underlying the difference between mGluR1a and 1b, we found that while the L1130stop construct showed the full Gs response, the T1125stop construct showed diminished Gs responses (> Figure 17‐5). There is a cluster of negatively charged amino acid residues (EEDE) situated between residues 1125 and 1130, and by mutating EEDE to QQQQ within fulllength mGluR1a, we were able to attenuate the Gs response. That cluster of negatively charged amino acid

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. Figure 17‐4 Switching among multiple signaling pathways governed by splice variation and by the presence of cytoplasmic-binding proteins. (a) Comparison of the functional coupling of mGluR1a (left) and mGluR1b (right) to signaling pathways mediated by Gq (changes [Ca2+]i, abscissa) and Gs (normalized [cAMP]i, ordinate). (b) Comparison of Gq (left) and Gs (right) signaling elicited by activation of mGluR1a and mGluR1b. Shown are means and SEMs. (c) Comparison of the functional coupling of mGluR1a, alone (left) and with homer1c (middle) or 4.1G (right), to signaling pathways mediated by Gq (changes [Ca2+]i, abscissa) and Gs (normalized [cAMP]i, ordinate). (d) Comparison of Gq (left) and Gs (right) signaling elicited by activation of mGluR1a, with or without binding proteins. Shown are means and SEMs. (Modified from original figures in Tateyama and Kubo (2007), with permission from Mol. Cell. Neurosci.)

residues thus appears to be critical for the coupling of mGluR1a to Gs and explains the difference between 1a and 1b, as this region is missing from mGluR1b (Tateyama and Kubo, 2007).

8 Factors Regulating the Switching Among Multiple Signaling Pathways 3—Effects of Cytoplasmic-Binding Protein We next focused on the effects of proteins known to bind to the C-terminal cytoplasmic region of mGluR1a. These include Homer1c, Homer1a (Tu et al., 1998; Abe et al., 2003a), tamalin (Kitano et al., 2002; Sugi et al., 2007), and 4.1G (Lu et al., 2004). When we coexpressed mGluR1a with Homer 1c in CHO cells, we observed little change in the Gs responses, whereas they were significantly weakened by coexpression of 4.1G (> Figure 17‐4c, d). To identify the structural determinant underlying this effect, we again examined the properties of a set of mGluR1a truncation mutants. We found that Gs signaling remained intact in a D1135stop mutant, and that it was inhibited by 4.1G. By contrast, when an L1130stop mutant was expressed, a clear Gs response was observed, even in the presence of 4.1G (> Figure 17‐5). By mutating a cluster of negatively charged amino acid residues (EEEED) located between residues 1130 and 1135 to QQQQQ within full-length mGluR1a, the suppression of Gs signaling by 4.1G was slightly but significantly

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. Figure 17‐5 Effects of truncation of the C-terminal tail on the coupling of mGluR1a to the Gs signaling pathway and on the inhibitory effect of 4.1G. (a) Coupling of full-length and truncated mGluR1a mutants to Gq (left, changes in [Ca2+]i) and Gs (right, normalized [cAMP]i) signaling in the absence (open columns) and presence (filled columns) of 4.1G. Shown are means and SEMs; * indicates statistically significant differences. (Modified from original figures in Tateyama and Kubo (2007), with permission from Mol. Cell. Neurosci.) (b) Region of the Cterminal cytoplasmic domain critical for coupling of mGluR1a to Gs signaling and for inhibition of that coupling by 4.1 G. EEDE and EEEED indicate clusters of negatively charged amino acid residues located between residues 1125–1130 and 1130–1135

reduced, verifying the importance of this region for the regulation by 4.1G. We also showed that the mGluR1a cytoplasmic region carrying the D1135stop mutation coimmunoprecipitated with 4.1G, but that with L1130stop mutation did not. It thus appears that 4.1G binds to wild-type mGluR1a at a cluster of negatively charged amino acid residues situated between residues 1130 and 1135, thereby suppressing the coupling of the receptor to Gs (Tateyama and Kubo, 2007).

9 Physiological Significance of Gd3+ Sensing Function In > Sections 4 and > 6, we showed that Gd3+ and glutamate bind to mGluR1 at different sites and that the resultant activated conformation and the downstream signaling pathways may also differ. However, the concentration of Gd3+ present in the cerebrospinal fluid is obviously not sufficient to activate mGluR1. What then is the physiological significance of the Gd3+ sensing function of mGluR1? One possibility is that it has no physiological significance, though the binding site is well designed and the effect is attractive from a biophysical point of view. An alternative possibility is that there is an, as yet unknown, endogenous molecule (substance X) present in the brain that interacts with mGluR1 at E238 instead of Gd3+, thereby facilitating transition to the Closed–Closed/Activated state. Suzuki et al. (2004) observed that negative cooperativity affecting glutamate binding to dimeric mGluR1 was diminished by 2-mM Cao2+, and suggested that the presence of Ca2+ at the dimer interface facilitates glutamate binding to the two subunits

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(i.e., the Closed/Closed conformation). Those findings suggest substance X is Cao2+. On the other hand, the ability of Cao2+ to enhance glutamate sensitivity was similar to wild-type mGluR1 and an E238Q mutant, suggesting Cao2+ is not substance X, which interacts with mGluR1 at E238 (Abe et al., 2003b). Polyamines, which are known to activate CaR (Quinn et al., 1997), are another strong candidate for substance X. These long thin molecules carry di-, tri-, or tetravalent positive charges and have a molecular size and shape suitable for binding to the slit at the dimer interface. We found that polyamines are unable to elicit mGluR1 activation (unpublished data), however, and suggest this possibility is unlikely. In summary, no clear candidate for substance X has yet been identified, and the search for substance X continues to be attractive.

10 Future Directions What will be the direction of research in the near future? (1) The importance of the conformational changes in the transmembrane domain in both class A and C receptors is now known (Binet et al., 2007). Detailed analysis of the intrasubunit conformational change in mGluR1, which we were unable to clearly detect using our initial construct (Tateyama et al., 2004), remains to be accomplished. (2) With respect to the regulation of switching among multiple pathways, mGluR1 is also known to couple to a Gi signaling pathway in some cells (Akam et al., 1997; Sharon et al., 1997). The regulatory mechanisms and relationship between Gq and Gi signaling will likely be a target of future study. (3) It also may be attractive and challenging to analyze the multiple signaling pathways at the single-molecule level by real-time optical recordings. (4) Toward clarification of the significance of the Gd3+ sensing function, a straightforward strategy would be to make mGluR1 knockin mice that carry an E238Q point mutation, which would not at all affect the sensitivity to glutamate or Cao2+ (Abe et al., 2003b). Any abnormality shown by these mutant mice could be interpreted to be the result of the loss of sensitivity to Gd3+ or substance X, confirming the physiological significance of this function. Finally, the dynamic aspects of the conformational changes and the functional regulation of this receptor will continue to be important research topics.

Acknowledgments The authors are supported in part by research grants from the Ministry of Education, Science, Sports, Culture and Technology of Japan (to Y.K. and to M.T.), from the Japan Science Promotion Society (to Y.K. and to M.T.), and from the Astellas Foundation for Research on Metabolic Disorders (to Y.K. and to M.T.).

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Akam EC, Carruthers AM, Nahorski SR, Challiss RAJ. 1997. Pharmacological characterization of type 1 alpha metabotropic glutamate receptor-stimulated [S-35]-GTP gamma S binding. Brit J Pharmacol 121: 1203-1209. Aramori I, Nakanishi S. 1992. Signal transduction and pharmacological characteristics of a metabotropic glutamate receptor, mGluR1, in transfected CHO cells. Neuron 8: 757-765. Baker JG, Hill SJ. 2007. Multiple GPCR conformations and signalling pathways: Implications for antagonist affinity estimates. Trends Pharmacol Sci 28: 374-381. Binet V, Duthey B, Lecaillon J, Vol C, Quoyer J, et al. 2007. Common structural requirements for heptahelical domain function in class A and class C G protein-coupled receptors. J Biol Chem 282: 12154-12163.

Structural rearrangement and functional regulation of the metabotropic glutamate receptor Brown EM, Gamba G, Riccardi D, Lombardi M, Butters R, et al. 1993. Cloning and characterization of an extracellular Ca(2+)-sensing receptor from bovine parathyroid. Nature 366: 575-580. De Blasi A, Conn PJ, Pin J, Nicoletti F. 2001. Molecular determinants of metabotropic glutamate receptor signaling. Trends Pharmacol Sci 22: 114-120. Hermans E, Challiss RA. 2001. Structural, signalling and regulatory properties of the group I metabotropic glutamate receptors: Prototypic family C G-protein-coupled receptors. Biochem J 359: 465-484. Houamed KM, Kuijper JL, Gilbert TL, Haldeman BA, O’Hara PJ, et al. 1991. Cloning, expression, and gene structure of a G protein-coupled glutamate receptor from rat brain. Science 252: 1318-1321. Jensen AA, Greenwood JR, Brauner-Osborne H. 2002. The dance of the clams: Twists and turns in the family C GPCR homodimer. Trends Pharmacol Sci 23: 491-493. Jensen AA, Sheppard PO, Jensen LB, O’Hara PJ, BraunerOsborne H. 2001. Construction of a high affinity zinc binding site in the metabotropic glutamate receptor mGluR1: Noncompetitive antagonism originating from the amino-terminal domain of a family C G-proteincoupled receptor. J Biol Chem 276: 10110-10118. Jingami H, Nakanishi S, Morikawa K. 2003. Structure of the metabotropic glutamate receptor. Curr Opin Neurobiol 13: 271-278. Joly C, Gomeza J, Brabet I, Curry K, Bockaert J, et al. 1995. Molecular, functional, and pharmacological characterization of the metabotropic glutamate receptor type 5 splice variants: Comparison with mGluR1. J Neurosci 15: 3970-3981. Kammermeier PJ, Yun J. 2005. Activation of metabotropic glutamate receptor 1 dimers requires glutamate binding in both subunits. J Pharmacol Exp Ther 312: 502-508. Kano M, Hashimoto K, Chen C, Abeliovich A, Aiba A, et al. 1995. Impaired synapse elimination during cerebellar development in PKC gamma mutant mice. Cell 83: 12231231. Kaupmann K, Huggel K, Heid J, Flor PJ, Bischoff S, et al. 1997. Expression cloning of GABA(B) receptors uncovers similarity to metabotropic glutamate receptors. Nature 386: 239-246. Kaupmann K, Malitschek B, Schuler V, Heid J, Froest W, et al. 1998. GABA(B)-receptor subtypes assemble into functional heteromeric complexes. Nature 396: 683-687. Kitano J, Kimura K, Yamazaki Y, Soda T, Shigemoto R, et al. 2002. Tamalin, a PDZ domain-containing protein, links a protein complex formation of group 1 metabotropic glutamate receptors and the guanine nucleotide exchange factor cytohesins. J Neurosci 22: 1280-1289.

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Kniazeff J, Bessis AS, Maurel D, Ansanay H, Prezeau L, et al. 2004. Closed state of both binding domains of homodimeric mGlu receptors is required for full activity. Nat Struct Mol Biol 11: 706-713. Kobilka BK, Deupi X. 2007. Conformational complexity of G-protein-coupled receptors. Trends Pharmacol Sci 28: 397-406. Kubo Y, Miyashita T, Murata Y. 1998. Structural basis for a Ca2+ -sensing function of the metabotropic glutamate receptors. Science 279: 1722-1725. Kubo Y, Tateyama M. 2005. Towards a view of functioning dimeric metabotropic receptors. Curr Opin Neurobiol 15: 289-295. Kubokawa K, Miyashita T, Nagasawa H, Kubo Y. 1996. Cloning and characterization of a bifunctional metabotropic receptor activated by both extracellular calcium and glutamate. FEBS Letters 392: 71-76. Kunishima N, Shimada Y, Tsuji Y, Sato T, Yamamoto M, et al. 2000. Structural basis of glutamate recognition by a dimeric metabotropic glutamate receptor. Nature 407: 971-977. Leach K, Sexton PM, Christopoulos A. 2007. Allosteric GPCR modulators: Taking advantage of permissive receptor pharmacology. Trends Pharmacol Sci 28: 382-389. Lu D, Yan H, Othman T, Rivkees SA. 2004. Cytoskeletal protein 4.1G is a binding partner of the metabotropic glutamate receptor subtype 1 alpha. J Neurosci Res 78: 49-55. Masu M, Tanabe Y, Tsuchida K, Shigemoto R, Nakanishi S. 1991. Sequence and expression of a metabotropic glutamate receptor. Nature 349: 760-765. Mitri C, Parmentier ML, Pin JP, Bockaert J, Grau Y. 2004. Divergent evolution in metabotropic glutamate receptors. A new receptor activated by an endogenous ligand different from glutamate in insects. J Biol Chem 279: 9313-9320. Miyashita T, Kubo Y. 2000a. Extracellular Ca2+ sensitivity of mGluR1 alpha associated with persistent glutamate response in transfected CHO cellsRecept Channels 7: 25-40. Miyashita T, Kubo Y. 2000b. Extracellular Ca2+ sensitivity of mGluR1 alpha induces an increase in the basal cAMP level by direct coupling with Gs protein in transfected CHO cell Recept Channels 7: 77-91 Muto T, Tsuchiya D, Morikawa K, Jingami H. 2007. Structures of the extracellular regions of the group II/III metabotropic glutamate receptors. Proc Natl Acad Sci USA 104: 37593764. Nakanishi S, Masu M, Bessho Y, Nakajima Y, Hayashi Y, et al. 1994. Molecular diversity of glutamate receptors and their physiological functions. EXS 71: 71-80. Nash MS, Saunders R, Young KW, Challiss RA, Nahorski SR. 2001. Reassessment of the Ca2+ sensing property of a type I metabotropic glutamate receptor by simultaneous measurement of inositol 1,4,5-trisphosphate and Ca2+ in single cells. J Biol Chem 276: 19286-19293.

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Parnot C, Kobilka B. 2004. Toward understanding GPCR dimers. Nat Struct Mol Biol 11: 691-692. Pin JP, Galvez T, Prezeau L. 2003. Evolution, structure, and activation mechanism of family 3/C G-protein-coupled receptors. Pharmacol Ther 98: 325-354. Pin JP, Kniazeff J, Goudet C, Bessis AS, Liu J, et al. 2004. The activation mechanism of class-C G-protein coupled receptors. Biol Cell 96: 335-342. Pin JP, Kniazeff J, Liu J, Binet V, Goudet C, et al. 2005. Allosteric functioning of dimeric class C G-proteincoupled receptors. FEBS J 272: 2947-2955. Quinn SJ, Ye CP, Diaz R, Kifor O, Bai M, et al. 1997. The Ca2+-sensing receptor: A target for polyamines. Am J Physiol 273: C1315-C1323. Rondard P, Liu J, Huang S, Malhaire F, Vol C, et al. 2006. Coupling of agonist binding to effector domain activation in metabotropic glutamate-like receptors. J Biol Chem 281: 24653-24661. Saunders R, Nahorski SR, Challiss RA. 1998. A modulatory effect of extracellular Ca2+ on type 1alpha metabotropic glutamate receptor-mediated signalling. Neuropharmacology 37: 273-276. Selkirk JV, Price GW, Nahorski SR, Challiss RA. 2001. Cell type-specific differences in the coupling of recombinant mGlu1alpha receptors to endogenous G protein subpopulations. Neuropharmacology 40: 645-656. Sharon D, Vorobiov D, Dascal N. 1997. Positive and negative coupling of the metabotropic glutamate receptors to a G protein-activated K+ channel, GIRK, in Xenopus oocytes. J Gen Physiol 109: 477-490. Sugi T, Oyama T, Muto T, Nakanishi S, Morikawa K, et al. 2007. Crystal structures of autoinhibitory PDZ domain of Tamalin: Implications for metabotropic glutamate receptor trafficking regulation. EMBO J 26: 2192-2205.

Sugiyama H, Ito I, Hirono C. 1987. A new type of glutamate receptor linked to inositol phospholipid metabolism. Nature 325: 531-533. Suzuki Y, Moriyoshi E, Tsuchiya D, Jingami H. 2004. Negative cooperativity of glutamate binding in the dimeric metabotropic glutamate receptor subtype 1. J Biol Chem 279: 35526-35534. Tabata T, Aiba A, Kano M. 2002. Extracellular calcium controls the dynamic range of neuronal metabotropic glutamate receptor responses. Mol Cell Neurosci 20: 56-68. Tateyama M, Abe H, Nakata H, Saito O, Kubo Y. 2004. Ligand-induced rearrangement of the dimeric metabotropic glutamate receptor 1alpha. Nat Struct Mol Biol 11: 637-642. Tateyama M, Kubo Y. 2006. Dual signaling is differentially activated by different active states of the metabotropic glutamate receptor 1alpha. Proc Natl Acad Sci USA 103: 1124-1128. Tateyama M, Kubo Y. 2007. Coupling profile of the metabotropic glutamate receptor 1alpha is regulated by the C-terminal domain. Mol Cell Neurosci 34: 445-452. Tsuchiya D, Kunishima N, Kamiya N, Jingami H, Morikawa K. 2002. Structural views of the ligand-binding cores of a metabotropic glutamate receptor complexed with an antagonist and both glutamate and Gd3+. Proc Natl Acad Sci USA 99: 2660-2665. Tu JC, Xiao B, Yuan JP, Lanahan AA, Leoffert K, et al. 1998. Homer binds a novel proline-rich motif and links group 1 metabotropic glutamate receptors with IP3 receptors. Neuron 21: 717-726. Zaccolo M, De Giorgi F, Cho CY, Feng L, Knapp T, et al. 2000. A genetically encoded, fluorescent indicator for cyclic AMP in living cells. Nat Cell Biol 2: 25-29.

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1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 346 2 AMPA Receptor Genes, Proteins, and Posttranslational Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . 346 3 Structure and Assembly of AMPA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 348 4 Channel Properties of AMPA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 348 5 AMPA Receptor Auxiliary Subunits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 349 6 Synaptic Localization of AMPA Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 351 7 Trafficking of AMPA Receptors to Synapses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 352 8 Roles of AMPA Receptor Interacting Proteins in AMPA Receptor Trafficking . . . . . . . . . . . . . . . . . . . 353 9 Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 354

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List of Abbreviations: ANQX, 6-azido-7-nitro-1,4-dihydroquinoxaline-2,3-dione; AMPA, amino-3hydroxy-5-methylisoxazole-4-propionic acid; CACNG1, Ca2+ channel gamma subunit 1; CACNG2, Ca2+ channel gamma subunit 2; CNQX, 6-cyano-7-nitroquinoxaline-2, 3-dione; EPSCs, excitatory postsynaptic currents; LTD, long-term depression; LTP, long-term potentiation; MAGUK, membrane-associated guanylate kinase; NMDA, N-methyl-D-aspartic acid; NP, neuronal pentraxin; NPR, NP receptor; NTD, N-terminal domain; PEPA, 4-[2-(phenylsulfonylamino)ethylthio]-2,6-difluoro-phenoxyacetamide; PSD, postsynaptic density; TARPs, transmembrane AMPA receptor regulatory proteins; UV, Ultraviolet

1 Introduction Information stored in the brain is encoded within neural circuits, which consist of billions of neurons. Neurons communicate with each other at synapses using neurotransmitters. Neuronal activity can modulate synaptic efficacy, which shapes the function of neural circuits. A major excitatory neurotransmitter in the brain is glutamate, and activity mediated by glutamate receptors modulates the neural circuits that underlie information storage in the brain. Glutamate released from presynaptic terminals binds to two types of glutamate receptors, metabotropic and ionotropic. Ionotropic glutamate receptors are further classified pharmacologically as AMPA- (amino-3-hydroxy5-methylisoxazole-4-propionic acid), NMDA- (N-methyl-D-aspartic acid), and kainate-sensitive glutamate receptors. These three receptor classes function distinctly in the brain. AMPA receptors play major roles in fast synaptic transmission, whereas NMDA and kainate receptors modulate synaptic efficacy. Due to their fast kinetics, AMPA receptors play major roles in controlling excitatory postsynaptic currents (EPSCs), and modulation of AMPA receptor activity at synapses results in alterations in neural circuits. In this chapter, I describe the ‘‘life of AMPA receptors,’’ from the expression of receptor subunits to the formation of functional receptors and their trafficking into and out of the synapses where they are active.

2 AMPA Receptor Genes, Proteins, and Posttranslational Modifications There are 16 known subunits of ionotropic glutamate receptors (GluR1–7, KA1–KA2, NR1, NR2A-D, NR3A-B), which are categorized by the type of functional receptor they are found in (AMPA, NMDA, or kainate). There are four AMPA receptor subunits, GluR1–GluR4. The first cDNA of a glutamate receptor, GluR-K1 (GluR1/A), was isolated as a kainate-gated ion channel using expression cloning in Xenopus laevis oocytes (Hollmann et al., 1989). Subsequently, GluR2–GluR4 (B–D) were isolated by DNA homology (Boulter et al., 1990; Keinanen et al., 1990; Nakanishi et al., 1990; Sakimura et al., 1990). Because GluR1–GluR4 display AMPA-selective pharmacology (AMPA > glutamate > kainate), GluR1–GluR4 are defined as AMPA-sensitive glutamate receptors. GluR5–GluR7 and KA1–KA2 were also isolated by DNA homology to GluR1–GluR4, but these receptors are classified as kainate receptors because of their distinct pharmacology (Seeburg, 1993; Holmann and Heinemann, 1994). The topology of AMPA receptors was determined by three different experiments that utilized the sequence similarity of AMPA to bacterial amino acid (glutamine)-binding proteins, N-glycosylation sites, and protease sensitivity. In the first experiment, two small parts of GluR (S1 and S2 in > Figure 18‐1) were found to resemble two small parts of bacterial amino acid-binding proteins, which form the ligand-binding domain and have been crystallized (Oh et al., 1993). Due to the location of S1 and S2 and the need for the ligand-binding domain to be exposed to the extracellular space, a topology of three transmembrane domains and one pore-loop was proposed for GluR1 (> Figure 18‐1). The extracellular domains of transmembrane proteins are often subjected to N-glycosylation at a consensus sequence, N-X-S/T. To identify which portions of GluR1 are extracellular, the consensus sequence for N-glycosylation was exogenously introduced into GluR1 in the second experiment. N-glycosylation was then used as a marker for extracellular location, and the results indicated that GluR1 has two extracellular domains (Hollmann et al., 1994). In the third experiment, Bennett and Dingledine (1995) used protease sensitivity experiments

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. Figure 18‐1 AMPA receptors and the TARP auxiliary subunit. AMPA receptors consist of a N-terminal domain (NTD), ligand-binding domain (S1S2), and three transmembrane domains with one pore-loop. The bound ligand is indicated by the gray circle marked ‘‘Glu’’ as Glutamate. The glutamine (Q) in the pore-loop is edited to an arginine residue (R) in the GluR2 subunit. A splicing site on the second extracellular loop generates the flip and flop isoforms. Narp and N-cadherin bind to the NTD, whereas several proteins bind to the cytoplasmic domain of AMPA receptors in a subunit (GluR1/GluR4 or GluR2/GluR3)-specific manner. The TARP auxiliary subunit has four membrane-spanning domains and one N-glycosylation site on the first extracellular loop

to show that both ends of the pore-loop are in the cytoplasm. Together, these three experiments support the presence of three transmembrane domains (TM1, TM2, and TM3), two extracellular domains, and one pore-loop (> Figure 18‐1). In the pore-loop of the GluR2 subunit, there is an RNA editing site where an RNA editing enzyme, ADAR2, deaminates adenosine and converts adenosine to inosine (Higuchi et al., 2000). As a result, an arginine residue is found in GluR2 even though the genomic DNA of all four isoforms encodes a glutamine codon (CAG) (Sommer et al., 1991). Several other residues in GluR2 are also edited. Importantly, GluR2 homomeric channels show linear current–voltage (I–V) relationships, but homomeric channels of GluR1, GluR3, and GluR4 show doubly rectifying I–V curves. This difference in ion permeability is defined by one amino acid (Q or R) in the pore-loop (Verdoorn et al., 1991). Because AMPA receptors in hippocampal pyramidal neurons show linear I–V relationships (Patneau and Mayer, 1991), functional hetero-oligomeric AMPA receptors in these neurons must contain at least one GluR2. Each of the 4 AMPA receptor subunits has 2 splicing isoforms involving 38 amino acids of the second extracellular loop, the so-called flip and flop isoforms, each of which contributes to distinct channel properties (see > Section 4) and distinct expression patterns in the brain and during development (Sommer et al., 1990; Monyer et al., 1991; Mosbacher et al., 1994). In addition, the cytoplasmic domains of GluR2 and GluR4 are also alternatively spliced (Kohler et al., 1994). Each AMPA receptor subunit in the brain is differentially phosphorylated, which modulates the properties and trafficking of the AMPA receptors. GluR1 and GluR4 have relatively longer cytoplasmic domains than GluR2 and GluR3 (Shi et al., 2001). GluR2 has PKC phosphorylation sites, and GluR1 and GluR4 have four sites phosphorylated by either PKA, PKC, or CaMKII (Roche et al., 1996; Barria et al., 1997; Mammen et al., 1997; Boehm et al., 2006) and one tyrosine phosphorylation site (Hayashi et al., 2000). These phosphorylation sites are proposed to modulate trafficking (see > Section 8).

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AMPA receptors are also palmitoylated at two cysteine residues in the pore-loop and the cytoplasmic domain. Palmitoylation of AMPA receptors has also been proposed to affect the trafficking of AMPA receptors (Hayashi et al., 2005).

3 Structure and Assembly of AMPA Receptors GluR1–GluR4 form hetero-oligomers in the brain (Wenthold et al., 1992; Brose et al., 1994). A series of experiments has identified the stoichiometry of the subunits of AMPA receptors. AMPA receptors show multiple concentration-dependent conductance states, and analysis of this behavior for both recombinant and native AMPA receptors suggests that AMPA receptors are tetrameric (Rosenmund et al., 1998; Smith and Howe, 2000). The N-terminal domain (NTD) of AMPA receptors can form homo- and heteromeric dimers with other AMPA receptor subunits and can also dimerize with kainate receptors, but with lower affinity (Leuschner and Hoch, 1999). The ligand-binding domain (S1S2) can form a dimer, and this dimerization is enhanced by the addition of a desensitization blocker, cyclothiazide (Sun et al., 2002). Native AMPA receptors purified from the rat brain appeared to have twofold symmetry in a single particle analysis at 30A˚ to 40A˚ resolution (Nakagawa et al., 2005), and the outer pore of AMPA receptors was also shown to have twofold symmetry (Sobolevsky et al., 2004). These results strongly support that the AMPA receptor tetramer is a dimer of dimers (> Figure 18‐2). The mechanisms for assembling AMPA receptor subunits have been examined. Characterization of the channel properties of chimeric AMPA/kainate receptors show that the second extracellular loop (S2) is necessary for dimer formation (Ayalon and Stern-Bach, 2001), whereas a residue in S1 is responsible for desensitization (Stern-Bach et al., 1998). The preferred subunit stoichiometry of heteromeric dimers was shown with a calcium permeability (Q/R) mutant in recombinant systems to be two GluR1 subunits and two GluR2 subunits (Mansour et al., 2001). Independently, Q/R editing was proposed to mediate AMPA receptor tetramerization (Greger et al., 2003). Biochemical experiments of overexpressed receptors in neurons show that GluR2(R) forms a dimer, whereas GluR2(Q) forms a tetramer. Along with dimerization of NTD, the Q/R editing site might be a mechanism for assembly of GluR2-containing AMPA receptors. The structure of the ligand-binding domain (S1S2) has been studied extensively with various agonists at atomic resolution. Glutamate and kainate activate the AMPA receptor channel completely and partially, respectively. An X-ray crystal structure showed that binding of glutamate and kainate induces closure of the S1S2 angle by 20 and 12 , respectively (Armstrong et al., 1998; Armstrong and Gouaux, 2000). Because the degree of closure in S1S2 correlates with the size of the unitary currents evoked by these agonists (Jin et al., 2003), it seems that the closure of the S1S2 domain determines the channel opening. It would be interesting to see how changes in the ligand-binding domain induce the channel pore domains to open. Structural analyses of full-length AMPA receptors at atomic resolution will be required to examine the mechanisms further.

4 Channel Properties of AMPA Receptors AMPA receptors open upon agonist binding, followed by channel closure upon agonist removal (deactivation) or channel closure with continuous agonist binding (desensitization). The kinetics of each AMPA receptor subunit has been extensively characterized. AMPA receptors GluR1–GluR4 open within 1 ms, deactivate at about 3 ms, and desensitize at about 6 ms. The kinetics vary about two to fourfold among the subunits. The rise time of miniature EPSCs in neurons is almost identical to the rise time of glutamate-evoked currents from recombinant AMPA receptors. There is no rule to predict the kinetics of a heteromer of AMPA receptor subunits, but in most cases AMPA receptors containing flop isoforms show faster kinetics than those containing flip isoforms (Mosbacher et al., 1994; Erreger et al., 2004). The desensitization of AMPA receptors can be blocked by drugs, the so-called AMPA receptor potentiators (AMPAkines). AMPAkines, including cyclothiazide, PEPA (4-[2-(phenylsulfonylamino) ethylthio]-2,6-difluoro-phenoxyacetamide), aniracetam, and TCM (trichlormethiazide) show isoform

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. Figure 18‐2 Assembly of AMPA receptor subunits. The proposed mechanism for tetramer formation is that the N-terminal domains (NTDs) of AMPA receptor subunits form dimers, and subsequently the dimers assemble with each other through the ligand-binding domain (S1S2) and transmembrane domain (TMD) to form tetramers (dimer of dimers)

specificity to either flip or flop. Structural and functional data support the idea that channel desensitization is linked to destabilization of the AMPA receptor dimer (Sun et al., 2002; Robert et al., 2005; Armstrong et al., 2006). Interestingly, AMPAkines have been reported to enhance cognitive function and are currently being investigated as potential treatments for a variety of neurological disorders, including schizophrenia, Alzheimer’s disease, and Parkinson’s disease (Staubli et al., 1994; Lynch et al., 1997; Li et al., 2003; O’Neill et al., 2004).

5 AMPA Receptor Auxiliary Subunits In some neurons, kainate evoked steady-state currents of AMPA receptors are 100-fold larger than those evoked by glutamate, whereas this is not the case in recombinant AMPA receptors expressed in heterologous cells (> Figure 18‐3). This discrepancy was reconciled when a group of AMPA receptor auxiliary proteins, the stargazin-like transmembrane AMPA receptor regulatory proteins (TARPs), was found (Nicoll et al., 2006; Ziff, 2007). TARPs consist of five isoforms, TARP g-2/stargazin, g-3, g-4, g-7, and g-8 (Tomita et al., 2003; Kato et al., 2007), and are evolutionally conserved (Walker et al., 2006).

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. Figure 18‐3 The addition of TARPs cause the channel properties of recombinant AMPA receptors to resemble those of native receptors. (a) AMPA receptors expressed in Xenopus laevis oocytes responded to glutamate (Glu) better than kainate (KA). (b) AMPA receptors co-expressed with TARP in Xenopus laevis oocytes responded to kainate better than glutamate. (c) Neuronal AMPA receptors in hippocampal CA1 pyramidal cells responded to kainate better than glutamate, which is similar to the recombinant AMPA receptors co-expressed with TARP

The prototypical TARP, g-2/stargazin, was identified as the causative gene of the stargazer and waggler phenotypes in mice. Both of these strains were isolated at the Jackson Laboratory as spontaneous mutants showing ataxia and absence epilepsy (Noebels et al., 1990; Letts et al., 1998). Originally, g-2/stargazin was thought to be identified as a Ca2+ channel gamma subunit 2 (CACNG2) because of its sequence similarity to Ca2+ channel gamma subunit 1 (CACNG1) (Letts et al., 1998). Subsequently, six other homologous genes (CACNG3–CACNG8) were identified (Burgess et al., 2001; Chu et al., 2001). All eight proteins have four transmembrane domains, and four of them (stargazin/g-2, 3, 4, and 8) have a typical class I PDZ domain-binding motif (-TTPV) at the C terminus (> Figure 18‐1). A key breakthrough in understanding the role of TARPs came from the discovery that stargazer and waggler show a loss of AMPA receptor activity at the mossy fiber–granule cell synapses in the cerebellum, but not in the hippocampus (Chen et al., 1999; Hashimoto et al., 1999). The reduction in AMPA receptor activity is caused by a deficit of AMPA receptor trafficking to the cell surface in stargazer cerebellar granule cells (Chen et al., 2000). Interestingly, overexpression of stargazin lacking the last four amino acids in cultured stargazer cerebellar granule cells rescues AMPA receptor activity at the cell surface, but not at synapses (Chen et al., 2000), which led to a two-step model of AMPA receptor trafficking. In this model, AMPA receptors are trafficked to the cell surface by stargazin and localize a synapses through the last four amino acids of stargazin, which interact with PSD95-like membraneassociated guanylate kinase (MAGUK) proteins through PDZ domains (Chen et al., 2000; Schnell et al., 2002; Dakoji et al., 2003). Each TARP isoform is expressed distinctly in the brain (Tomita et al., 2003; Fukaya et al., 2005). Stargazer mice, which lack the cerebellum-enriched TARP g-2/stargazin, show no AMPA receptor activity at cerebellar granule cell synapses, whereas mice lacking the hippocampus-abundant TARP g-8 show a loss of surface AMPA receptor activity and a 30% reduction in the amplitude of AMPA receptor-mediated EPSCs in the hippocampus (Rouach et al., 2005; Fukaya et al., 2006). Furthermore, mice with disruptions in both stargazin and g-8 show greater reductions in AMPA receptor-mediated EPSCs than stargazer or g-8 single-knockout mice (Rouach et al., 2005; Fukaya et al., 2006), suggesting that there is functional redundancy among the TARP family proteins. How do TARPs regulate AMPA receptor activity? TARPs co-immunoprecipitate only with AMPA receptor GluR1–GluR4 from detergent-soluble brain extracts (Tomita et al., 2004). TARPs form a complex of over 600 kDa on native PAGE, which overlaps in size with the AMPA receptor-containing complexes

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(Vandenberghe et al., 2005). Purification of AMPA receptors from transgenic mouse or rat brain also showed TARPs to be a major component of the AMPA receptor complex (Fukata et al., 2005; Nakagawa et al., 2005). TARPs increase AMPA receptor activity in Xenopus laevis oocytes (Chen et al., 2003). In addition to modulating trafficking, TARPs also modulate the channel properties of AMPA receptors (Tomita et al., 2005a). TARPs slow the desensitization and deactivation of AMPA receptors (Priel et al., 2005; Tomita et al., 2005a) and decrease the EC50 of glutamate slightly (Yamazaki et al., 2004; Priel et al., 2005; Tomita et al., 2005a) by increasing the rate of channel opening, as shown by a single channel analysis (Tomita et al., 2005a). Surprisingly, TARPs modulate the response of AMPA receptors to kainate (> Figure 18‐3). As mentioned in > Section 4, native AMPA receptors respond to kainate better than glutamate, whereas recombinant AMPA receptors do not. Co-expression of stargazin with AMPA receptors robustly increases kainate-evoked steady-state currents in Xenopus laevis oocytes, indicating that neuronal AMPA receptors interact with TARPs (Tomita et al., 2005a; Turetsky et al., 2005; Kott et al., 2007). Further analysis revealed that stargazin modulates AMPA receptor trafficking through its cytoplasmic domain, and AMPA receptor channel properties through its first extracellular loop (Tomita et al., 2005a).

6 Synaptic Localization of AMPA Receptors A characteristic structure of excitatory synapses in the brain is the postsynaptic density (PSD), and PSD95 has been identified as major components of the PSD (Cho et al., 1992). Subsequently, its homologous proteins have been identified and grouped as MAGUKs. Because AMPA- and NMDA-type glutamate receptors function at the PSD, a molecular link between glutamate receptors and PSD95 was expected. PSD95 directly binds specifically to NMDA receptors, but not AMPA receptors, through the PDZ domain (Kornau et al., 1995). Unexpectedly, overexpression of PSD95 in cultured hippocampal neurons increased AMPA receptor-mediated EPSCs, but not NMDA receptor-mediated EPSCs (El-Husseini et al., 2000). Furthermore, acute knockdown of PSD95-like MAGUKs by RNAi specifically decreased AMPA receptor-mediated EPSCs (Elias et al., 2006). However, PSD95 knockout mice had no obvious change in AMPA receptor-mediated EPSCs (Migaud et al., 1998), probably because other MAGUK isoforms, including PSD93, SAP102, and SAP97, compensate for the chronic loss of PSD95. The analysis of mice with multiple isoforms knocked out will clarify functional redundancy of MAGUKs. Because PSD95 cannot bind directly to AMPA receptors, there might be an intermediate molecule that can interact with both AMPA receptors and PSD95-like MAGUKs. Among the candidates for such intermediate molecules are the TARPs (Dakoji et al., 2003; Tomita et al., 2003). Overexpression of TARP g-2/stargazin restores AMPA receptor-mediated EPSCs in stargazer cerebellar granule cells that have no AMPA receptor activity, whereas overexpression of stargazin lacking the last four amino acids, which is the binding domain for PSD95-like MAGUKs, does not (Chen et al., 2000). These results clearly show that TARPs are involved in the synaptic localization of AMPA receptors. However, it remains unclear whether all AMPA receptors at synapses interact with TARPs (TARPin AMPA receptor) or whether some AMPA receptors do not have TARPs (TARPless AMPA receptors). It is also not clear whether TARP-dependent synaptic localization of AMPA receptors depends on neuronal types or brain regions. Further analysis is necessary to clarify these issues. Proteins of the neuronal pentraxin (NP) family promote synaptic clustering of AMPA receptors (O’Brien et al., 1999; Sia et al., 2007). The NP family consists of two secreted isoforms, NP1 and NP2/Narp, and one membrane-spanning isoform, NP receptor (NPR). These three isoforms form hetero-oligomers in the brain (Xu et al., 2003). Interestingly, Narp (neuronal activity regulated protein) was originally isolated as an immediate early gene whose transcription is induced by neuronal activity. Overexpression of recombinant Narp increases the numbers of excitatory synapses, but not the number of inhibitory synapses. Furthermore, Narp induces clustering of AMPA receptors in receptor-transfected HEK cells (O’Brien et al., 1999). NP1 interacts with the NTD of GluR4, and this interaction is sufficient to induce GluR4 clustering in GluR4-transfected heterologous cells cocultured with neurons (Sia et al., 2007).

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Conversely, NP1 RNAi knockdown decreases the clustering of GluR4 in interneurons (Sia et al., 2007). Both studies showed that NP binds to the extracellular domain of AMPA receptors (O’Brien et al., 1999; Sia et al., 2007).

7 Trafficking of AMPA Receptors to Synapses AMPA receptors move to synapses to function as neurotransmitter receptors. There are two types of trafficking in general, vesicular trafficking and lateral diffusion. However, it remains uncertain whether AMPA receptors are first expressed on the cell surface and then move into synapses by lateral diffusion or are trafficked in vesicles and exocytosed at synapses (> Figure 18‐4a). Biochemical and cell biological studies have shown that AMPA receptors can be trafficked by both vesicular trafficking and lateral diffusion (> Figure 18‐4b). To analyze the kinetics of exo- and endocytosis of neuronal proteins, Ehlers labeled cell surface proteins with biotin in neuronal cultures, incubated the cultures at 37 C, and quantified the amount of intracellular biotintylated proteins (Ehlers, 2000). This study showed that the endo- and exocytosis of AMPA receptors are regulated by neuronal activity. Importantly, NMDA induced the recycling of AMPA receptors, whereas AMPA induced the endocytosis of AMPA receptors into lysosomes for degradation. Several groups independently used imaging techniques to visualize AMPA- or NMDA-induced endocytosis of AMPA receptors (Carroll et al., 1999; Beattie et al., 2000; Lin et al., 2000; Passafaro et al., 2003). Imaging of GFP-tagged clathrin in living neurons showed

. Figure 18‐4 Trafficking of AMPA receptors to synapses. (a) Possible pathways by which AMPA receptors are trafficked to synapses are shown. (1) to synapses by vesicular trafficking from the trans-Golgi network (TGN); (2) to the surface of the soma by exocytosis and then to synapses by lateral diffusion; (3) to the surface of the soma, followed by endocytosis at the soma or dendrites and then vesicular trafficking to synapses. (b) Possible pathways of synaptic insertion of AMPA receptors on spines. AMPA receptors could insert into extrasynaptic sites by lateral diffusion (1) or exocytosis (4). AMPA receptors move to the postsynaptic density (PSD) by lateral diffusion (2) or exocytosis (5). For removal from synapses, AMPA receptors are endocytosed from synapses (6), move to the extrasynaptic region by lateral diffusion (3), followed by endocytosis (7)

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that there is a hot spot for clathrin-mediated endocytosis next to the PSD (7 in > Figure 18‐4b) (Blanpied et al., 2002). AMPA receptors can be inserted into the plasma membrane and move into synapses by lateral diffusion (Borgdorff and Choquet, 2002). Neuronal activity increases the rate of lateral diffusion into synapses (Groc et al., 2004), and PSD95 stabilizes laterally diffusing AMPA receptors around synapses (Bats et al., 2007). Interestingly, the rate of lateral diffusion can also be regulated by synaptic activity (2 in > Figure 18‐4b) (Ehlers et al., 2007). Lateral diffusion is also supported by electrophysiological experiments with a new drug called ANQX (6-azido-7-nitro-1,4-dihydroquinoxaline-2,3-dione) (Adesnik et al., 2005). ANQX, a derivative of the competitive antagonist CNQX (6-cyano-7-nitroquinoxaline-2, 3-dione), is a photoreactive, irreversible antagonist of AMPA receptors (England, 2006). By illuminating a restricted area of an ANQX-treated neuron with ultraviolet (UV) light, the dynamics of AMPA receptors at synapses and the cell surfaces of neuronal cell bodies can be observed. Surprisingly, in ANQX-treated neurons in which lateral diffusion is blocked by UV illumination, the recovery of AMPA receptor activity took longer than 12 h (Adesnik et al., 2005). Because 12 h seem too long for active synaptic trafficking of AMPA receptors, this result suggests that lateral diffusion plays a major role in the synaptic trafficking of AMPA receptors, although there is a possibility that ANQX may silence whole neurons by depleting surface AMPA receptor activity. N-cadherin binds to the NTD of AMPA receptor subunit GluR2 to regulate the lateral diffusion of AMPA receptors (> Figure 18‐1). Overexpression of GluR2 increases spine size and density in hippocampal pyramidal cells and interneurons. This enhancement is mediated by the NTD of GluR2, but not GluR1 or GluR3 (Passafaro et al., 2003). A proteomic analysis of GluR2 using primary cultured neurons treated with the cell-impermeable cross linker DTSSP identified several proteins, including N-cadherin, as interacting with GluR2 (Saglietti et al., 2007). N-cadherin-coated beads recruit GluR2 in cultured hippocampal neurons. Furthermore, knockdown of N-cadherin prevents changes in GluR2-induced spine morphology (Saglietti et al., 2007). Because the NTD is known to be involved in the dimerization of AMPA receptors (Leuschner and Hoch, 1999), it will be necessary to further investigate the mechanisms of the interaction of NTD with N-cadherin.

8 Roles of AMPA Receptor Interacting Proteins in AMPA Receptor Trafficking The NMDA receptor-mediated modulation of synaptic strength known as long-term potentiation (LTP) is mediated by changes in the number of AMPA receptors at synapses (Manabe et al., 1992). Several proteins have been identified as AMPA receptor interacting proteins that modulate AMPA receptor trafficking, which may help to understand the mechanisms regulating synaptic insertion of AMPA receptors. The cytoplasmic domains of AMPA receptors are classified into two groups, long tailed (GluR1 and GluR4) and short tailed (GluR2 and R3). The long-tailed subunits contain a type I PDZ domainbinding motif (A-T/S-GL) at the C terminus, whereas the short-tailed subunits have a type II PDZ domain-binding motif (SVKI) at the C terminus (Songyang et al., 1997; Shi et al., 2001). The PDZ domains of AMPA receptor interacting proteins bind highly selectively to either short- or long-tailed AMPA receptors and are proposed to have distinct functions. SAP97 interacts with GluR1 through the PDZ domain (Leonard et al., 1998) and is suggested to function in localization or trafficking in the ER (Sans et al., 2001). However, no obvious difference was observed in SAP97-disrupted mice or mice lacking the last seven amino acids of GluR1 (Klocker et al., 2002; Kim et al., 2005). Band 4.1 was isolated as a GluR1/GluR4-binding protein that promotes the surface expression of GluR1/GluR4 receptors (Shen et al., 2000; Coleman et al., 2003). Because the homologous protein 4.1 in erythrocytes is involved in cytoskeletal structure, Band 4.1 may connect glutamate receptors to the cytoskeleton. More proteins have been isolated that bind to the short-tailed GluR2 and R3. PDZ domains in GRIP/ABP and PICK1 recognize the C-terminal four amino acids, -SVKI, of GluR2/3. The serine residue in the last four amino acids of GluR2 is phosphorylated in the brain and by PKC in vitro (McDonald et al., 2001). GRIP/ABP recognizes only the nonphosphorylated form of GluR2, whereas PICK1 recognizes both the nonphosphorylated and phosphorylated forms of GluR2 (Chung et al., 2000). This binding specificity

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suggests that GRIP/ABP and PICK1 have distinct roles. Cerebellar long-term depression (LTD) is mediated by PKC downstream of metabotropic GluR1 (mGluR1; Linden and Connor, 1991; Aiba et al., 1994). Both PICK1 knockout mice and GluR2 K882A knockin mice, which cannot be phosphorylated by PKC, show no cerebellar LTD (Steinberg et al., 2006). These results suggest a model in which, upon phosphorylation of GluR2 at synapses by PKC, PICK1 binds to phosphorylated GluR2 and removes GluR2-containing receptors from synapses. Two proteins involved in the endocytotic machinery, AP2 and NSF, have also been identified as interacting with GluR2/3. AP2 and NSF bind to the middle of the cytoplasmic domain of GluR2/3 (> Figure 18‐1). Inhibiting the interaction of NSF with GluR2 by peptide injection decreases basal synaptic transmission, but LTD is still observed. Conversely, inhibition of AP2 by peptide injection does not change basal synaptic transmission, but prevents LTD (Lee et al., 2002), suggesting that AP2 and NSF have distinct roles in basal and activity-dependent trafficking of AMPA receptors. However, GluR2/3 double-knockout mice show enhanced LTD in the hippocampus, suggesting that LTD is mediated not only by AP2-dependent endocytosis, but also by other unidentified mechanisms. TARPs also contribute to AMPA receptor trafficking. Stargazin-disrupted stargazer mice show no AMPA receptors at the surfaces of cerebellar granule cells and increased amounts of the immature nonN-glycosylated form of AMPA receptors in neurons (Chen et al., 2000; Tomita et al., 2003). Overexpression of stargazin restores AMPA receptor activity in stargazer cerebellar granule cells lacking endogenous stargazin (Chen et al., 2000). However, overexpression of stargazin in wild-type hippocampal neurons increases AMPA receptor activity only at the cell surface, and not at synapses (Schnell et al., 2002). This result suggests that there is another modulator for regulating the synaptic insertion of AMPA receptors. Interestingly, TARPs are highly phosphorylated in the brain (Tomita et al., 2005b; Inamura et al., 2006). TARP g-2/stargazin has nine phosphorylation sites in its cytoplasmic domain in cultured neurons, and overexpression of a stargazin mutant protein that mimics the constitutively phosphorylated state increases synaptic AMPA receptor activity twofold, whereas mutants proteins that mimic the nonphosphorylated state do not increase activity. Furthermore, experiments in neurons expressing various stargazin phosphorylation mutants have also shown that phosphorylated residues are required for the expression of LTP and LTD (Tomita et al., 2005b). The phenotypes of mice transgenic at these sites are interesting to study possible roles of TARP phosphorylation in animal behaviors.

9 Concluding Remarks AMPA receptors play major roles in controlling the excitatory synaptic strength that underlies many aspects of brain functions. Because synaptic plasticity requires the regulation of AMPA receptor activity, extensive studies have been done to understand the molecular mechanisms that regulate AMPA receptor activity and remarkable progress has been made. However, many fundamental issues remain unresolved. How do AMPA receptors assemble in the brain? AMPA receptors are very likely to be a dimer of dimers. However, details, including the assembly domains and roles of TARP auxiliary subunits, remain uncertain. A dominant model is that the initial dimerization occurs through the NTDs, and a dimer of NTD dimers is subsequently formed through the ligand-binding domain (S1S2) or the transmembrane domains (> Figure 18‐2). However, AMPA receptors lacking the NTD still form functional channels. Furthermore, all AMPA receptors in cerebellar granule cells contain TARPs. How do TARPs affect the assembly of AMPA receptors? How many TARPs incorporate into one AMPA receptor? The structure of AMPA receptors at atomic levels will be required to fully understand AMPA receptor assembly. The simplest model for synaptic localization of AMPA receptors is that TARPs bridge AMPA receptors to PSD95-like MAGUKs. Although all AMPA receptors in the cerebellar granule cells contain TARPs, it remains uncertain whether this is true for all AMPA receptors in other brain regions. AMPA receptors, TARPs, and MAGUKs all have four isoforms, which are expressed distinctly in the brain. There could therefore be isoform specificity for assembly or localization. In addition, roles have been proposed for the cytoplasmic domain of AMPA receptors, especially in GluR2, on the basis of experiments using overexpressed proteins. However, GluR2/GluR3 double-knockout mice have synaptic AMPA receptors.

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Furthermore, GluR1 localizes at synapses, but GluR1 knockout mice show no changes in EPSCs. These complicated phenomena may suggest the presence of compensation mechanisms like homeostatic plasticity to adjust synaptic strength. Do AMPA receptors contain their own trafficking signal? Studies with overexpressed AMPA receptor mutants suggested that the AMPA receptor trafficking signal is in the cytoplasmic domain. However, genetic studies with AMPA receptor knockout or knockin mice did not fully support this model, as described in the previous paragraph. How can we be convinced? A signal or protein should be both necessary and sufficient for synaptic AMPA receptor localization and trafficking. Distinct roles of different AMPA receptor cytoplasmic domains are sufficient but not necessary, because GluR2/GluR3 knockout mice still show both basal transmission- and activity-dependent modulation. TARP-mediated synaptic localization is necessary but not sufficient, because overexpression of wild-type TARP does not increase synaptic AMPA receptors, although there may be other limiting factors like the number of MAGUKs or the TARP phosphorylation states. There could be multiple independent mechanisms for synaptic localization and trafficking of AMPA receptors in response to environmental changes. Additional studies are needed to generate general models to explain both synaptic localization and trafficking of AMPA receptors and to further classify the types of AMPA receptors at synapses.

Acknowledgments I thank James R. Howe and members of the Tomita lab for helpful discussions and Hillel Adesnik for providing > Figure 18‐3c. ST is supported by the Esther A. & Joseph Klingenstein Fund, the Edward Mallinckrodt Jr. Foundation, and NARSAD.

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P2 Purinergic Receptor

K. Inoue

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 362

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P2 Purinergic Receptor in the Central Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 362

3 3.1 3.1.1 3.1.2

Structure and Function of P2X Family in Neural Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 P2X4 and Pain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364 The Mechanism of Microglial P2X4-Dependent Neuropathic Pain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365 The Mechanism of P2X4R Upregulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367

4 4.1 4.2 4.3

Structure and Function of P2Y Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367 P2Y1 and Glio-Transmission in Astrocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 368 P2Y12 and Microglial Chemotaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 369 P2Y6 and Microglial Phagocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 370

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Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371

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2009 Springer Science+Business Media, LLC.

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Abstract: Accumulating findings indicate that interactions between extracellular nucleotides and P2 purinergic receptors play important roles in cell-to-cell communication in the central nervous system (CNS), even though ATP is recognized primarily to be a source of free energy and nucleotides are key molecules in cells. P2 purinoceptors are divided into two families, ionotropic receptors (P2X family) and metabotropic receptors (P2Y family). P2X receptors (seven types; P2X1–P2X7) contain intrinsic pores that open by binding with ATP. P2Y receptors (eight types; P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13, and P2Y14) are activated by nucleotides and couple to intracellular second-messenger systems through heteromeric G-proteins. Nucleotides are released or leaked from nonexcitable cells as well as neurons in physiological and pathophysiological conditions. Glia is the interesting nonexcitable cell and is classified into astrocytes, oligodendrocytes, and microglia. Astrocytes express many types of P2 purinoceptors and release the ‘‘gliotransmitter’’ ATP to communicate with neurons, microglia, and the vascular walls of capillaries. Microglia also express many types of P2 purinoceptors and are known as resident macrophages in the CNS. ATP and other nucleotides work as ‘‘warning molecules,’’ especially through activating microglia in pathophysiological conditions. Microglia play a key role in neuropathic pain and show phagocytosis through nucleotide-evoked activation of P2X4 and P2Y6 receptors, respectively. Molecular-, cellular-, and system-level evidences for extracellular nucleotide signaling place P2 purinergic receptors in the central stage in the function of CNS. List of Abbreviations: abmeATP, a,b-methylene ATP; 2-MeSADP, 2-methylthio-ADP; adBDNF, BDNFtransducing recombinant adenovirus; anti-TrkB, antibody against the TrkB receptor; BDNF siRNA, short interfering RNA directed against BDNF; CNS, central nervous system; ERK, extracellular signal regulated protein kinase; IP3, inositol trisphosphate; KA, kainic acid; LI, lamina I; MAP, mitogen-activated protein; NMDA, N-methyl-D-aspartate; PI3K, phosphatidylinositol 3-kinase; PS, phosphatidylserine; PTX, pertussis toxin; TNP-ATP, 20 ,30 -O-(2,4,6-trinitrophenyl)-ATP; TrkB–Fc, BDNF-sequestering fusion protein; UTP, uridine 50 -triphosphate

1

Introduction

In 1972, the role of intracellular ATP was already established as the source of free energy to maintain life, and intracellular nucleotides were known as key molecules within cells. With such times background, Burnstock (1972) coined new roles of extracellular nucleotides as neurotransmitters in the pond. This action, of course, caused tsunamis and the idea had to meet with severe criticism from ‘‘so-called scientists.’’ In 1993, however, P2Y1, a subtype of cell-surface nucleotide receptors called P2 purinoceptors, was cloned (Webb et al., 1993). Afterward, numerous subtypes of these receptors were cloned, which led scientists to gradually accept the notion of a ‘‘purinergic nervous system’’ (Burnstock and Knight, 2004). Now, P2 purinoceptors are divided into two families, ionotropic P2X receptor family and metabotropic P2Y receptor family (> Figure 19-1). Recently, accumulating evidences suggest that nucleotides are released or leaked from nonexcitable cells as well as neurons and play a role in cell-to-cell communication in physiological and pathophysiological conditions of CNS. One of the most interesting cell type in nonexcitable cells is glia that make up over 70% of the total cell population in CNS, and are classified into astrocytes, oligodendrocytes, and microglia. Astrocytes express many types of P2 purinoceptors and release ATP spontaneously or in response to various stimuli, and communicate with neurons at synapses, and microglial cells and vascular walls at capillaries. Microglia also express many types of P2 purinoceptors and are known as resident macrophages in CNS, accounting for 5–10% of the total population of glia (Kreutzberg, 1996; Stoll and Jander, 1999). According to the recent evidences indicating the importance of P2 purinergic receptors, many paragraphs of this chapter are allotted to the functions of P2 purinergic receptors in the glia.

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P2 Purinergic Receptor in the Central Nervous System

P2 purinergic receptors are divided into two families, ionotropic P2X family and metabotropic P2Y family (> Figure 19-1). P2X receptors (seven types; P2X1–P2X7) contain intrinsic pores allowing ions that switch

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. Figure 19‐1 P2 purinergic receptors. P2 purinergic receptors are divided into two families, ionotropic P2X family and metabotropic P2Y family. P2X family (seven types; P2X1–P2X7) has two transmembrane domains. P2X receptors consist of three subunits and contain intrinsic pores allowing ions to flow that switch conformation from closed to open on binding ATP. P2Y (eight types; P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13, and P2Y14) are activated by extracellular purine, pyrimidine nucleotides or sugar-nucleotides and are coupled to intracellular secondmessenger systems through heteromeric G-proteins. Basically, P2Y1, P2Y2, P2Y4, P2Y6, P2Y11 couple to Gq/11, and P2Y12, P2Y13, and P2Y14 couple to Gi/o. P2Y11 also couples to Gs

conformation from closed to open on binding ATP to flow. P2Y (eight types; P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13, and P2Y14) is activated by extracellular purine, pyrimidine nucleotides, or sugarnucleotides and couples to intracellular second-messenger systems through heteromeric G-proteins (Burnstock and Knight, 2004). Both families of P2 purinergic receptors are expressed in whole body including the central nervous system (CNS). In CNS, these receptors are expressed in neurons, microglia, astrocytes, and oligodendrocytes, and are linked to signal transduction in these cell types through various manners depending on receptor subtypes.

3

Structure and Function of P2X Family in Neural Signaling

The P2X family has a variety of amino acid length ranging from 388 (shortest in P2X4) to 595 (longest in P2X7) in human. Each family has two hydrophobic, putative transmembrane segments (TM1 and TM2) that sandwich a large extracellular domain with ten conserved cysteine residues forming disulphide bonds. The amino (N-) and carboxy (C-) termini are intracellular. This topology resembles the form of epithelial Na channels and acid-sensing channels. It was reported that a P2X receptor/channel consists of three subunits (one subunit means one subtype of P2X family), and both TM1 and TM2 of one subunit may contribute to the pore, suggesting that six helices from three subunits could form a permeation pore. Most P2X receptors/channels are a nonselective cation channel through which Na+, Ca2+, and K+ penetrate. P2X5 receptor is significantly chloride permeable. Some P2X receptors also show significant permeability to divalent cations, with Ca2+ fluxes greater than nicotinic family and similar to those found in N-methyl-Daspartate (NMDA) receptors. Three molecules of ATP seem to bind to the extracellular portions of a P2X receptor (three subunits), that is, one molecule of ATP binds to one subunit.

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It appears that all three subunits are either identical (homomeric receptors) or not identical (heteromeric receptors). Lewis et al. (1995) first showed the ability of P2X3 to form heteromeric receptors with P2X2 (P2X2+3). Biochemical approaches using coimmunoprecipitation have shown that all P2X subunits except P2X7 can heteropolymerize with any of the other P2X subunits (Radford et al., 1997; Torres et al., 1999). The coassembly of P2X1+5 (Torres et al., 1998), P2X2+3 (Lewis et al., 1995), P2X2+6 (King et al., 2000), and P2X4+6 (Le et al., 1998) heteromeric receptors has been demonstrated to form functional receptors. In addition, there is a recent report that the P2X2+3 receptor seems likely to contain one P2X2 and two P2X3 subunits (Jiang et al., 2003). Each of the seven homomeric P2X receptors and at least four heteromeric receptors show different electrophysiological and pharmacological properties in terms of current kinetics, desensitization rates, and sensitivities to agonists and antagonists (Ralevic and Burnstock, 1998; Khakh et al., 2001; North, 2002). a,b-Methylene ATP (abmeATP), an analogue of ATP, is a useful agonist for basically identifying P2X receptors containing P2X1 or P2X3 subunit (i.e., P2X1, P2X3, P2X1+5, and P2X2+3), although abmeATP also activates P2X4+6 (Ralevic and Burnstock, 1998; Khakh et al., 2001; North, 2002). Suramin and PPADS have an antagonistic property to almost all P2X receptors except the rat P2X4 receptor (Buell et al., 1996). 20 ,30 -O-(2,4,6-trinitrophenyl)-ATP (TNP-ATP) at a nanomolar range selectively blocks P2X1, P2X3, and P2X2+3 receptors (Virginio et al., 1998) and, thus, would be a useful tool to distinguish those from the other P2X receptors that are about 1,000 times less sensitive to TNP-ATP (Virginio et al., 1998). Furthermore, diinosine pentaphosphate (Ip5I) is a very potent antagonist to P2X1 (King et al., 1999) and P2X3 receptors but, interestingly, not to P2X2 and P2X2+3 receptors (Dunn et al., 2001). This is the only antagonist for distinguishing between P2X3 and P2X2+3. Jarvis et al. (2002) recently developed the first selective antagonist for P2X3 (and P2X2+3), A-317491. A-317491 blocks the responses mediated by P2X3 or P2X2+3 receptors in a competitive fashion without any effect on other receptors, enzymes, and ion channels (Jarvis et al., 2002). The P2X family is widely expressed in neurons and glia. In neuronal communication, ATP is a fast neurotransmitter where it may be released with other transmitters at some synapses in CNS. ATP-evoked synaptic currents in the postsynaptic neuron are small, and measured in a subpopulation of neurons. The functional significance at the postsynapses might be related to the high Ca2+ fluxes through P2X receptors, even at resting membrane potentials. The Ca2+ entry during ATP synaptic transmission affects the frequency dependence for the induction of synaptic plasticity. P2X2 and P2X4 subunits are found in excitatory synapses in CNS, however, they exist at the periphery of the postsynaptic density. This suggests that postsynaptic P2X2 and P2X4 receptors are only activated during bouts of action potential firing, when released ATP in synaptic cleft can spill out to activate these receptors. The physiological meanings of postsynaptic P2X family are still unclear. Presynaptic P2X channels play important roles in CNS. For example, in the hippocampus, presynaptic P2X actions form one component of ATP effects in the circuit formed among CA3 pyramidal neurons, GABAergic interneurons, and CA1 pyramidal neurons. The stimulation of presynaptic P2X2 receptors of CA3 neurons increase the release of glutamate onto GABAergic interneurons, whereas postsynaptic P2Y1 receptors depolarize interneurons and astrocytic P2Y receptors evoke glutamate release onto interneurons. The net effect on output neurons is dominated by GABAergic synaptic inhibition because there are few preor postsynaptic ATP receptors on CA1 pyramidal neurons. In the nucleus of the solitary tract, activation of presynaptic P2X channels evoked large-amplitude miniature synaptic currents, suggesting that Ca2+ entry through presynaptic P2X channels may trigger multivesicular release, or release of glutamate from an otherwise ‘‘silent’’ set of vesicles or terminals.

3.1 P2X4 and Pain P2X4 subunits have recently been implicated in pain sensation. In this case, it seems that the critical sites of P2X4 expression are spinal microglia rather than neurons. The pain in response to noxious stimuli has an important role as an early warning device that alerts us to the presence of damaging stimuli. This normal ‘‘good’’ pain usually goes away after the nociceptive stimulus is removed or the tissue damage is repaired. There is, however, a type of pain that does not go away even though the tissue has already healed. One type

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of this pain is called neuropathic pain, which typically develops when pain-related neurons are damaged through surgery, bone compression in cancer, diabetes, or infection. Neuropathic pain is a major factor causing impaired quality of life in more than 15 million people worldwide and, unfortunately, is frequently resistant to all known drugs. We are now beginning to understand that neuropathic pain is not just a symptom of disease but is a consequence of disordered functioning of the nervous system (Woolf and Mannion, 1999; Woolf and Salter, 2000; Scholz and Woolf, 2002). Over the last decade, accumulating evidence concerning how peripheral nerve injury creates neuropathic pain has suggested that nerve injury produces molecular and cellular alterations that result in multiple forms of neuronal plasticity and anatomical reorganization in the dorsal horn of the spinal cord. These alterations have been proposed to be crucial in the pathogenesis of neuropathic pain (Woolf and Mannion, 1999; Woolf and Salter, 2000; Scholz and Woolf, 2002). Although the dominant theme in research on neuropathic pain has been to understand the roles of neurons in the peripheral nervous system and the dorsal horn, there is a rapidly growing body of evidence indicating spinal glial cells, in particular microglia, play a critical role in the pathogenesis of neuropathic pain. Tsuda et al. (2003) directly implicated activated microglia in the pathogenesis of neuropathic pain by determining the role of the purinoceptor P2X4 receptor (P2X4R). A clue to identifying P2X4Rs in the spinal cord as being required for neuropathic pain first came from a pharmacological investigation of pain behavior after nerve injury using antagonists (Tsuda et al., 2003). The marked tactile allodynia after injury of a spinal nerve was found to be reversed by administering intrathecally TNP-ATP (blocking P2X4) but was unaffected by PPADS (not blocking P2X4).Thus, it was inferred that the tactile allodynia depends upon P2X4Rs in the spinal cord. The expression of P2X4R protein progressively increased in the days following nerve injury, the time course of which was parallel to that of the development of tactile allodynia. In the immunohistochemical analysis, it was found that activated microglia in the dorsal horn of the nerve-injured side were positive for P2X4R protein. Moreover, it was found that reducing the upregulation of P2X4R protein in spinal microglia by P2X4R antisense oligodeoxynucleotide prevented the development of the tactile allodynia. The sufficiency of P2X4R activation in microglia for the development of allodynia was demonstrated by intrathecal administration of cultured microglia stimulated in vitro by ATP. In naive animals, allodynia develops progressively over a period of 3–5 h following the administration of P2X4R-stimulated microglia (Tsuda et al., 2003). Moreover, in rats in which tactile allodynia was caused by the ATP-stimulated microglia, this allodynia was reversed by administering TNP-ATP. The allodynia caused by ATP-stimulated microglia is pharmacologically similar to that caused by peripheral nerve injury. These findings indicate that P2X4R stimulation of microglia is not only necessary for tactile allodynia but is also sufficient to cause allodynia (> Figure 19-2).

3.1.1 The Mechanism of Microglial P2X4-Dependent Neuropathic Pain It was already reported that the nerve injury-induced tactile allodynia depends on a depolarizing shift in the Eanion of spinal lamina I (LI) neurons in the dorsal spinal cord, resulting in converting the GABAA-receptorand glycine-receptor-mediated inhibition to excitation (Coull et al., 2003). This report caused an idea that microglia may affect Eanion in LI neurons. To investigate this possibility, microglia were administered to the lumbar spinal level of naive rats via an intrathecal catheter as described (Tsuda et al., 2003). Administering microglia stimulated with ATP caused a progressive tactile allodynia over the 5 h after injection. Voltage-clamp recording was made from LI neurons of slices prepared 5 h after intrathecal microglia administration. Eanion in LI neurons from rats administered ATP-stimulated microglia was shifted to 61.6 mV from 68.3 mV, that is, Eanion in spinal slices taken from normal rats (Coull et al., 2005). In addition, using current clamp recordings, GABA response switched from hyperpolarizing in control rats to depolarizing in rats treated with ATP-stimulated microglia. Activated microglia secrete various biologically active molecules, including BDNF, which was implicated in the hypersensitivity of dorsal horn neurons in sensitization and inflammation (Mannion et al., 1999; Thompson et al., 1999; Heppenstall and Lewin, 2001) and in anion gradient shifts in the hippocampus (Rivera et al., 2002).

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. Figure 19‐2 Mechanism how P2X4 receptors in activated microglia evoke neuropathic pain in the dorsal horn. Activated microglia in the spinal cord after nerve injury express P2X4 receptors. P2X4R activation leads to the release of bioactive diffusible factors such as BDNF and cytokines. BDNF causes a collapse of transmembrane anion gradient in postsynaptic neurons of dorsal horn lamina I presumably through the downregulation of KCC2. This changes the action of GABA and glycine into excitatory from inhibitory in postsynaptic neurons. These neurons are innervated by inhibitory interneurons that release GABA and glycine after the depolarizing stimulation of glutamate from Ab dorsal root ganglion neurons. Thus, touch sensation may cause strong pain. The net hyperexcitability in the dorsal horn pain network by these factors from activated microglia may be responsible for neuropathic pain

To examine whether BDNF could trigger shifts in pain hypersensitivity and in LI neuronal Eanion similar to those resulting from the application of ATP-stimulated microglia, recombinant BDNF was administered intrathecally to normal rats. BDNF produced tactile allodynia comparable with that produced by ATPstimulated microglia. Eanion of LI neurons in slices treated with BDNF showed a depolarizing shift. By perfusion with BDNF, the proportion of neurons responding to GABA with a rise in [Ca2+]i increase reached 31% of neurons recorded between 80 and 120 min. The rise in [Ca2+]i was prevented by the GABAA receptor blocker bicuculline, confirming that the effect was mediated by GABAA receptors. Thus, acute administration of BDNF in slices caused a depolarizing shift in Eanion and caused GABA to produce net excitation. To examine the effects of prolonged exposure to BDNF in vivo, a BDNF-transducing recombinant adenovirus (adBDNF) (Gravel et al., 1997) was administered intrathecally to the rats. A progressive tactile allodynia was observed over the 4 days after the treatment of adBDNF. Eanion in LI neurons from adBDNFinjected rats was significantly less negative than that from control rats. Thus, similar to acute administration of BDNF, sustained local release of BDNF caused the allodynia and a depolarizing shift in Eanion. Moreover, a function-blocking antibody against the TrkB receptor (anti-TrkB) and a BDNF-sequestering fusion protein (TrkB–Fc) acutely inhibited the allodynia and the shift of Eanion of LI neurons. These findings

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indicate that endogenous BDNF is necessary to sustain both the tactile allodynia and the depolarizing shift in Eanion in LI neurons that result from nerve injury (Coull et al., 2005). The administration of ATP-stimulated microglia with either anti-TrkB or TrkB–Fc did not develop tactile allodynia. After pretreatment of microglia with double-stranded short interfering RNA directed against BDNF (BDNF siRNA), the ATP-stimulated microglia injected intrathecally into normal rats did not cause the allodynia. Anti-TrkB and BDNF siRNA prevented the shift in Eanion induced by ATP-stimulated microglia. ATP stimulation caused release of BDNF from microglia in culture. This effect of ATP was blocked by treating the cultures with the P2X receptor blocker TNP-ATP. In addition, pretreatment of the microglia with BDNF siRNA prevented release of BDNF by ATP stimulation. By bath-application of TNPATP to spinal slices taken from allodynic rats 2 weeks after nerve injury, Eanion of LI neurons returned to normal value. Thus, P2X4 receptor activation is necessary to sustain the depolarizing shift in Eanion in rats with nerve injury (Coull et al., 2005). These findings show that both the decrease in paw withdrawal threshold and the shift in Eanion in LI neurons caused by ATP-stimulated microglia through P2X4 require BDNF–TrkB signaling (> Figure 19-2).

3.1.2 The Mechanism of P2X4R Upregulation The upregulation of P2X4R in microglia is an important process in producing neuropathic pain. Nasu-Tada et al. (2006) reported the role of fibronectin, an extracellular matrix protein, as a potential candidate molecule in the P2X4R upregulation in microglia. It was found that microglia cultured on fibronectincoated dishes showed a marked increase in P2X4R expression both at the mRNA and protein levels. Intrathecal delivery of ATP-stimulated microglia to the rat lumber spinal cord revealed that microglia treated with fibronectin more effectively induced allodynia than control microglia (Nasu-Tada et al., 2006). Whether fibronectin is involved in the upregulation of P2X4Rs in spinal microglia after peripheral nerve injury in vivo is yet to be determined. It was also reported that activating both TLRs and NOD2 (another pattern-recognition receptor) in cultured microglia increased the expression of P2X4R at the mRNA level (Guo et al., 2006), thus suggesting the involvement of these receptors in the regulation of P2X4R.

4

Structure and Function of P2Y Family

P2Y receptors can be broadly subdivided pharmacologically into four groups. First group receptors including human and rodent P2Y1, P2Y12, and P2Y13, and human P2Y11 respond to adenine nucleotides like ADP and ATP. Second group receptors including human P2Y4 and P2Y6 respond to uracil nucleotides like uridine 50 -triphosphate (UTP) or UDP. Third group receptors including human and rodent P2Y2, rodent P2Y4 and P2Y11 have mixed selectivity to adenine and uracil nucleotides. Forth group receptor (P2Y14) responds solely to the sugar nucleotides like UDP-glucose and UDP-galactose. P2Y receptor genes do not contain introns in the coding sequence, except for the P2Y11 receptor. The experiment using sitedirected mutagenesis of the P2Y1 and P2Y2 receptors suggests that some positively charged residues in TM 3, 6, and 7 are crucial for binding of nucleotides (Abbracchio et al., 2006). These residues may interact with the negative charges of the phosphate groups of nucleotides. All P2Y receptors have an H-X-X-R/K motif in TM6. The P2Y1, P2Y2, P2Y4, P2Y6, and P2Y11 receptors have a Y-Q/K-X-X-R motif in TM7. Also P2Y12, P2Y13, and P2Y14 receptors have another motif, K-E-X-X-L (Abbracchio et al., 2006). P2Y receptors couple to various G proteins (> Figure 19-1). Indirect evidence was obtained from several cellular responses, that is, intracellular Ca2+, inositol phosphates, or cAMP concentration with or without pertussis toxin (PTX) sensitivity. Direct evidence was obtained by measuring the effect of ADP on GTP hydrolysis in vesicles reconstituted with P2Y receptors and G proteins. P2Y1 couples to either Gaqb1g2 or Ga11b1g2 (Waldo and Harden, 2004). P2Y2 couples to PLC-b1 via Gaq/11 and to PLC-b3 via Gai3b1g2derived bg subunits (Murthy and Makhlouf, 1998). P2Y12 couples to Gai2 more effectively than to Gai1 and Gai3, but not to Gao or Gaq (Bodor et al., 2003). When P2Y11 receptor is activated by ATP, it leads to a rise in cAMP and in inositol trisphosphate (IP3) and cytosolic calcium, whereas activation by

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UTP of P2Y11 leads to calcium mobilization without IP3 or cAMP increase (White et al., 2003). The P2Y12 receptor activates phosphatidylinositol 30 -kinase (PI3K) via Gai, but also RhoA and Rho kinase (Soulet et al., 2004), and Nasu-Tada et al. (2005) recently reported that a P2Y12-mediated decrease in cyclic AMP is an important step in membrane ruffling and chemotaxis by microglia on fibronectin-coated dishes (Nasu-Tada et al., 2005). The P2Y13 receptor can couple to G16, Gi, and Gs at high concentrations of ADP. Three signaling pathways are characterized by different ratios of ADP to 2-methylthio-ADP (2-MeSADP) potency. These data suggest an interesting evidence that when a subtype of P2Y receptor couples to distinct several G proteins, distinct active conformations of the receptor may provide differently agonist-specific signaling pathways (Marteau et al., 2003). The activation of several P2Y receptors is commonly associated with the stimulation of several mitogen-activated protein (MAP) kinases, in particular extracellular signal regulated protein kinase (ERK) 1/2. According to the cell types and the P2Y receptor subtypes, other classes of the MAP kinases, PKC, calcium, and PI3-K are found to be involved to a variable extent (Abbracchio et al., 2006).

4.1 P2Y1 and Glio-Transmission in Astrocytes Astrocytes express various neurotransmitter receptors including metabotropic glutamate receptors, dopamine receptors, noradrenalin receptors, serotonin receptors, and P2 receptors (Haydon, 2001). After the first reports of elevations in [Ca2+]i in individual cultured astrocytes in response to neurotransmitters (Enkvist et al., 1989; Cornell-Bell et al., 1990), it became apparent that many neurotransmitters stimulate Ca2+ elevations through specific receptors in glial cells. Cornell-Bell et al. (1990) showed that glutamate can elicit [Ca2+]i not only in individual cells, but also intercellular waves of increased [Ca2+]i that are propagated from single cells to multiple neighboring cells (Cornell-Bell et al., 1990). Dani et al. (1992) showed that neuronal activity can directly initiate such Ca2+ waves in astrocytes networks (Dani et al., 1992). Other stimuli such as local mechanical or electrical stimulation were subsequently observed to initiate similar intercellular Ca2+ signaling in astrocytes. For some years, such Ca2+ waves had been thought to propagate via gap junctions (Boitano et al., 1992; Sneyd et al., 1994; Sneyd et al., 1995), through which inositol 1,4,5-trisphosphate (InsP3) can be diffused to mobilize Ca2+ release (Sneyd et al., 1995). More recently, it was reported that Ca2+ waves can be propagated between astrocytes, even when the cells do not contact each other directly, and the extent and direction of the Ca2+ wave propagation are significantly influenced by movement of the extracellular medium (Guthrie et al., 1999). Now, it is suggested that the extracellular molecule ATP could be a primary signal for the Ca2+ wave propagation and important in cross talk among astrocytes and even other cell types in the CNS. This means that one of ‘‘gliotransmitters’’ is extracellular ATP. Actually, ATP is released from astrocytes during Ca2+ wave propagation (Guthrie et al., 1999; Cotrina et al., 2000), and the propagation can be reduced or abolished by a purinergic antagonists (Guthrie et al., 1999; Fam et al., 2000; Koizumi et al., 2003) or the ATP-degrading enzyme apyrase (Guthrie et al., 1999; Koizumi et al., 2003). In addition, visualization of the release of ATP demonstrated that the velocity of ATP release well correlates with that of the Ca2+ wave in astrocytes (Koizumi et al., 2003). So far, the physiological significance of the ATP-evoked increase in [Ca2+]i in astrocytes themselves has received only limited attention. The stimulation of astrocytes with ATP enhances mitogenic signaling via the ERKmediated pathway, increases proliferation (Bolego et al., 1997), and protects astrocytes against oxidative stress (Shinozaki et al., 2004). Further comprehensive studies will reveal the importance of the ATPmediated Ca2+ responses in astrocytes. Neurons express a wide variety of P2X and P2Y receptor subtypes in the pre- and postsynaptic regions, and ATP directly mediates synaptic transmission as a fast neurotransmitter in the rat medial habenula (Edwards et al., 1992) and in the spinal cord dorsal horn (Bardoni et al., 1997). In addition, exogenously applied ATP potentiates (Gu and MacDermott, 1997; Hugel and Schlichter, 2000; Kato and Shigetomi, 2001; Shigetomi and Kato, 2004) or inhibits (Koizumi and Inoue, 1997; Cunha and Ribeiro, 2000) synaptic transmission in the CNS. Given that astrocytic Ca2+ waves can evoke changes in neuronal synaptic activity and are mediated by the release of ATP, ATP released from astrocytes should be involved in astrocyteto-neuron signaling in synaptic regions of the CNS. As described earlier, astrocytes lack the ability to

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propagate regenerative electrical signals but are nonetheless responsive to a variety of extracellular stimuli and produce regenerative Ca2+ waves that spread within astrocyte networks (Guthrie et al., 1999; Fam et al., 2000; Newman, 2001; Koizumi et al., 2003), for which gliotransmitter ATP has a central role. Thus, gliotransmitter ATP would affect neuronal activity, especially synaptic transmission. ATP from astrocytes has recently been shown to decrease the excitability of neurons in the retina (Newman, 2003) and mediate presynaptic inhibition in cultured hippocampal neurons (Koizumi et al., 2003; Zhang et al., 2003). Cultured hippocampal neurons reveal synchronous spontaneous Ca2+ oscillation, which is extracellular Ca2+ dependent, tetrodotoxin sensitive, and inhibited by inhibitors of ionotropic glutamate receptors, suggesting that the neuronal Ca2+ oscillation is mediated by glutamatergic synaptic transmission (Ogura et al., 1987; Koizumi and Inoue, 1997; Koizumi et al., 2003). ATP released from astrocytes downregulated the spontaneous neuronal Ca2+ oscillation (Koizumi et al., 2003) and EPSCs in a hippocampal culture (Zhang et al., 2003) by inhibiting presynaptic functions of glutamatergic neurons. Similarly, astrocytic ATP-mediated presynaptic inhibition was observed in the hippocampus in situ (Zhang et al., 2003) although adenosine, a metabolite of ATP degraded by ectonucleotidases, also functioned as an inhibitory molecule in the slices. Recently, Pascual et al. (2005) demonstrated that, in transgenic mice that express a dominantnegative SNARE domain selectively in astrocytes, the release of transmitters from these glial cells was blocked. By releasing ATP, which accumulates as adenosine, astrocytes tonically suppress synaptic transmission. These results indicate that astrocytes are intricately linked in the regulation of synaptic strength and plasticity and provide a pathway for synaptic cross talk (Pascual et al., 2005). It is hypothesized that glutamate and ATP from astrocytes evoke opposing actions to modulate excitatory and inhibitory synaptic transmission, respectively. In addition to mediating inhibitory rather than excitatory effects on synaptic transmission, ATP-mediated astrocyte-to-neuron signaling further differs from glutamatedependent signaling mechanisms by the fact that it occurs in a tonic fashion (Zhang et al., 2003; Koizumi et al., 2003; Pascual et al., 2005). Application of apyrase induces a potentiation of spontaneous neuronal Ca2+ oscillations or EPSCs in the absence of any astrocytic stimulation, suggesting the presence of a constitutive ATP-dependent inhibition of synaptic transmission. Furthermore, spontaneous astrocytic Ca2+ responses occur in both purified astrocyte cultures and mixed cultures of astrocytes and neurons. The spontaneous Ca2+ signals in astrocytes were inhibited by apyrase but persisted in the presence of TTX. These data suggest that astrocytes constitutively release ATP, which exerts tonic downregulation of excitatory synaptic transmission (Koizumi et al., 2003; Zhang et al., 2003).

4.2 P2Y12 and Microglial Chemotaxis Extracellular ATP works as a chemoattractant. Microglial chemotaxis by ATP via P2Y12 Rs was originally found by Honda et al. (2001), and was confirmed in vivo in P2Y12 R knockout animals (Haynes et al., 2006). Neuronal injury results in the release or leakage of ATP, which appears to be a ‘‘find-us’’ signal from damaged neurons to cause microglial chemotaxis (> Figure 19-3). P2Y12 acts as rapid sensor of extracellular nucleotide, which causes microglia chemotactic activity toward injured site in CNS (Haynes et al., 2006; Ohsawa et al., 2007). Of these subtypes, P2X4 and P2Y6 are upregulated after nerve injury, whereas P2Y12 is expressed from resting state and selectively in brain microglia but not in peripheral macrophages (Sasaki et al., 2003; Haynes et al., 2006). More recently, it was reported that the P2X4 receptor (P2X4R) is also involved in ATP-induced microglial chemotaxis in the intracellular signaling pathway downstream of P2Y12R (Ohsawa et al., 2007). Ohsawa et al. examined the effect of two PI3K inhibitors, wortmannin and LY294002, on chemotaxis in a Dunn chemotaxis chamber. The PI3K inhibitors significantly suppressed chemotaxis without affecting ATP-induced membrane ruffling. ATP stimulation increased Akt phosphorylation in the microglia, and the increase was reduced by the PI3K inhibitors and a P2Y12R antagonist. These results indicated that P2Y12R-mediated activation of the PI3K pathway is required for microglial chemotaxis in response to ATP. They also found that the Akt phosphorylation was reduced when extracellular calcium was chelated. This result was unexpected and suggested that ionotropic P2X receptors are involved in microglial chemotaxis by affecting the PI3K pathway. Therefore, they tested the effect of various P2X4R antagonists on the chemotaxis and found that pharmacological blockade of P2X4R significantly inhibited

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. Figure 19‐3 Microglial chemotaxis and phagocytosis stimulated by nucleotides. Damaged or over excited neurons in traumatic or ischemic injury leak or release a large amount of ATP and UTP. Microglia might be attracted by ATP/ADP and come near the damaged cells because the concentration of ATP is higher. Thus, ATP seems to be a ‘‘find-us’’ signal. Neuronal injury caused an increase in extracellular UTP, which was immediately metabolized into UDP. UDP is a diffusible molecule that signals the crisis of damaged neurons to microglia, triggering phagocytosis. Microglia subsequently recognize UDP, starting to recognize ‘‘eat-me’’ signals attached to the targets and engulf them. UDP is not likely a ‘‘eat-me’’ signal but ‘‘eat-us’’ signal

chemotaxis. In addition, knockdown of the P2X4R in microglia by RNA interference through the lentivirus vector system also markedly inhibited the ATP-induced microglial chemotaxis without affecting membrane ruffling, the same as the effect of PI3K inhibitors. These results indicated that the P2X4R-mediated calcium signaling may be involved in PI3K/Akt activation and regulate microglial chemotaxis. Local Ca2+ influx through P2X4R at membrane ruffles may be necessary for maintenance or enhancement of the local P2Y12R-activated PI3K signals (Ohsawa et al., 2007).

4.3 P2Y6 and Microglial Phagocytosis When neurons are injured or dead, microglia are activated, resulting in their interaction with immune cells, active migration to the site of injury, release of proinflammatory substances, and the phagocytosis of damaged cells or debris. For such activation of microglial motilities, extracellular nucleotides have a central role. In addition to microglial migration by ATP, another nucleotide, UDP, an endogenous agonist of the P2Y6 receptor, greatly activates the motility of microglia and orders microglia to eat damaged neurons. UDP does not cause chemotaxis, but instead causes phagocytosis by microglia (Koizumi et al., 2007). Phagocytosis, a specialized form of endocytosis, is the uptake by the cell of relatively large (>1.0 mm) particles into vacuoles and has a central role in tissue remodeling, inflammation, and the defense against infectious agents (Tjelle et al., 2000). Phagocytosis finally starts after the recognition of ‘‘eat-me’’ signal through various cell-surface phagocytosis receptors, including Fc receptors, complement receptors, integrins, endotoxin receptors (CD18, CD14), mannose receptors, and scavenger receptors (Lauber et al., 2003). Since recognition is the first and the most important step for phagocytosis, extensive studies on phagocytosis receptors have been reported. With regard to apoptotic cells, it is well known that dying cells express

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so-called ‘‘eat-me’’ signals such as phosphatidylserine (PS) on their surface membrane (Lauber et al., 2003), by which microglia recognize the apoptotic cells in order to catch and remove them (Lauber et al., 2003). As for amyloid b protein (Ab), a key molecule that mediates Alzheimer’s disease, microglia remove Ab presumably via Fc receptor-dependent phagocytosis (Schenk et al., 1999; Bard et al., 2000). It is worth to notice that generally speaking, these signals attach to the cell or debris. Therefore, it would seem to be difficult for phagocytes to recognize diffusible signals such as nucleotides. However, it was found that exogenous UDP caused microglial phagocytosis in a concentration-dependent manner, which was P2Y6 receptor dependent. Also, neuronal injury caused by kainic acid (KA) upregulated P2Y6 receptors in microglia, the KA-evoked neuronal injury resulted in an increase in extracellular UTP, which was immediately metabolized into UDP in vivo and in vitro, and UDP leaked from injured neurons caused P2Y6 receptor-dependent phagocytosis in vivo and in vitro (Koizumi et al., 2007). UDP could be a molecule that signals to microglia from damaged neurons suffering traumatic or ischemic injury. It should be noted that nucleotides could be both ‘‘find-us’’ and ‘‘eat-us’’ signals (> Figure 19-3). Cells release ATP, and we also found that KA caused an increase in extracellular UTP/UDP. Therefore, microglia might be attracted by ATP/ADP (Honda et al., 2001; Davalos et al., 2005; Nimmerjahn et al., 2005) and subsequently recognize UDP, leading to the removal of the dying cells and their debris. It is interesting that ATP/ADP is not able to efficiently activate P2Y6 receptors, nor can UDP act on P2Y12 receptors.

5

Conclusion

Brain is composed of the wide variety of cells that are neurons, astrocytes, microglia, vascular cells, and also invaded immune cells in some pathophysiological condition. Brain requires some means of intercellular communication that can integrate distinct elements into a dynamic and highly regulated function of brain. Extracellular ATP is a promising candidate for a key molecule in this communication, because all of these neural and nonneural cell types can release ATP for cell–cell communication, and all of these cells express some type of P2 purinergic receptors for extracellular ATP and its metabolites. This universality of the expression is quite an unique character of P2 purinergic receptors than other receptors. The second messenger systems activated by these receptors mediate a diverse range of nervous system processes, from the millisecond synaptic transmission to timescale responses. Recently, the function of P2 purinergic receptors in neuron–glia interaction attracts much attention in the neuroscience. Then, this chapter picked up reports of P2 purinergic receptors relating to pain, chemotaxis, and phagocytosis from huge amounts of important phenomenon reported in CNS. As already mentioned, P2 purinergic system seems to show the true value by playing a more important role in pathophysiological situation when they faced a crisis from normal time. To verify this context, we need more selective pharmacological tools and more accurate in vivo imaging methods. They will accelerate progress in discovering the broad scope of biological functions mediated by P2 purinergic signaling in neuron–glia interaction. In addition, new animal models are desired earnestly. Moreover, we have to verify the mechanisms for the release of nucleotides from several types of cells. For this purpose, we need new methods to probe released nucleotides. Using these fine tools, we will finally reach a goal to know the real functions of P2 purinergic systems in the brain.

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P2 purinergic receptor King BF, Townsend-Nicholson A, Wildman SS, Thomas T, Spyer KM, et al. 2000. Coexpression of rat P2X2 and P2X6 subunits in Xenopus oocytes. J Neurosci 20: 4871-4877. Koizumi S, Fujishita K, Tsuda M, Shigemoto-Mogami Y, Inoue K. 2003. Dynamic inhibition of excitatory synaptic transmission by astrocyte-derived ATP in hippocampal cultures. Proc Natl Acad Sci USA 100: 11023-11028. Koizumi S, Inoue K. 1997. Inhibition by ATP of calcium oscillations in rat cultured hippocampal neurones. Br J Pharmacol 122: 51-58. Koizumi S, Shigemoto-Mogami Y, Nasu-Tada K, Shinozaki Y, Ohsawa K, et al. 2007. UDP acting at P2Y6 receptors is a mediator of microglial phagocytosis. Nature 446: 1091-1095. Kreutzberg GW. 1996. Microglia: A sensor for pathological events in the CNS. Trends Neurosci 19: 312-318. Lauber K, Bohn E, Krober SM, Xiao YJ, Blumenthal SG, et al. 2003. Apoptotic cells induce migration of phagocytes via caspase-3-mediated release of a lipid attraction signal. Cell 113: 717-730. Le KT, Babinski K, Seguela P. 1998. Central P2X4 and P2X6 channel subunits coassemble into a novel heteromeric ATP receptor. J Neurosci 18: 7152-7159. Lewis C, Neidhart S, Holy C, North RA, Buell G, et al. 1995. Coexpression of P2X2 and P2X3 receptor subunits can account for ATP-gated currents in sensory neurons. Nature 377: 432-435. Mannion RJ, Costigan M, Decosterd I, Amaya F, Ma QP, et al. 1999. Neurotrophins: Peripherally and centrally acting modulators of tactile stimulus-induced inflammatory pain hypersensitivity. Proc Natl Acad Sci USA 96: 9385-9390. Marteau F, Le Poul E, Communi D, Communi D, Labouret C, et al. 2003. Pharmacological characterization of the human P2Y13 receptor. Mol Pharmacol 64: 104-112. Murthy KS, Makhlouf GM. 1998. Coexpression of ligandgated P2X and G protein-coupled P2Y receptors in smooth muscle. Preferential activation of P2Y receptors coupled to phospholipase C (PLC)-beta1 via Galphaq/11 and to PLC-beta3 via Gbetagammai3. J Biol Chem 273: 4695-4704. Nasu-Tada K, Koizumi S, Inoue K. 2005. Involvement of beta1 integrin in microglial chemotaxis and proliferation on fibronectin: Different regulations by ADP through PKA. Glia 52: 98-107. Nasu-Tada K, Koizumi S, Tsuda M, Kunifusa E, Inoue K. 2006. Possible involvement of increase in spinal fibronectin following peripheral nerve injury in upregulation of microglial P2X(4), a key molecule for mechanical allodynia. Glia 53: 769-775.

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Inhibitory Glycine Receptors

S. Dutertre . D. Kuzmin . B. Laube . H. Betz

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376

2 2.1 2.2 2.3 2.4

Overview of GlyR Structure: Subunit Composition, Ligand-Binding Sites, and Ion Channel Region . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376 Genes, Proteins, and Subunit Topology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376 Structure of the GlyR and Its Extracellular Ligand Binding Sites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Structure of the GlyR Transmembrane Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 Ion Selectivity and Drug-Binding Sites Within the Transmembrane Region . . . . . . . . . . . . . . . . . . 380

3 3.1 3.2 3.3 3.3.1 3.3.2 3.3.3 3.3.4 3.3.5

Expression and Functions of GlyR Subtypes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 381 GlyR Genes Display Distinct Expression Patterns in the Mammalian CNS . . . . . . . . . . . . . . . . . . . . 381 GlyRs Are Excitatory During Early Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 381 Roles of GlyR Isoforms in the Adult CNS: Lessons from Mutant Mice . . . . . . . . . . . . . . . . . . . . . . . . 382 a1 GlyRs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 382 a2 GlyRs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 382 a3 GlyRs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 383 a4 GlyRs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 383 GlyR b Subunit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 383

4 4.1

Human Disorders of Glycine Neurotransmission . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 384 Mutations in GlyR Genes Result in Hereditary Neuromotor Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . 384

5

Conclusion and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 384

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2009 Springer ScienceþBusiness Media, LLC.

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Abstract: The glycine receptor (GlyR) is a major inhibitory ligand-gated ion channel in the vertebrate central nervous system (CNS) that controls both motor and sensory pathways. This pentameric membrane protein is composed of α and β subunits that span the postsynaptic membrane and form a central anionselective channel. Molecular cloning has identified four genes coding for α subunits and one gene coding for the β subunit whose differential expression generates distinct GlyR subtypes in the mammalian CNS. Alternative splicing and RNA editing extend the heterogeneity of GlyR subunits and significantly complicate GlyR pharmacology. Recent structural insights from the X-ray structures of homologous receptor proteins complemented by modeling studies have allowed to interprete available mutational and pharmacological data and provided important clues to ligand binding and ion permeation mechanisms. GlyR mutations cause neuromotor diseases, and specific GlyR subtypes have been implicated in the regulation of pain perception. Hence, the rational design of GlyR active compounds might produce drugs with subtypeselective properties and therapeutic potential. This review summarizes recent data on the molecular organisation, ligand binding sites, ion channel region, expression and functions of GlyRs as well as on human disorders associated with defects in GlyR function and glycine neurotransmission. List of Abbreviations: GABA, g-aminobutyric acid; NMDA, N-methyl-D-aspartate; GABAA, gammaaminobutyric acid type-A; nAChR, nicotinic acetylcholine receptor; ECD, extracellular domain; AChBP, acetylcholine-binding protein; SCAM, substituted cysteine accessibility method; sIPSCs, spontaneous inhibitory postsynaptic currents; PGE2, prostaglandin E2; 5HT; receptor, serotonin type 3 receptor

1

Introduction

In order to function normally, the CNS requires complex mechanisms that integrate both excitatory and inhibitory signals. g-Aminobutyric acid (GABA) and glycine are the two major inhibitory neurotransmitters in the adult mammalian CNS. Both amino acids also act as excitatory neurotransmitters at embryonic neurons. In addition, glycine serves as a co-agonist at excitatory synapses, where it promotes the activation of N-methyl-D-aspartate (NMDA) receptors by glutamate. Glycinergic synapses mediate fast inhibitory neurotransmission mainly in spinal cord, brain stem, and caudal brain, and are involved in a variety of motor and sensory functions (Laube et al., 2002). Historically, the amino acid glycine was first suspected to act as a neurotransmitter when it was found in higher concentration in spinal cord as compared with other brain regions (Aprison and Werman, 1965). Its inhibitory role was demonstrated later by the reduced firing of spinal neurons following application of glycine (Werman et al., 1967). Glycine exerts its inhibitory effects by binding to specific receptors at the postsynaptic membrane, the inhibitory glycine receptors (GlyRs). Binding of glycine leads to the opening of the GlyR’s anionic channel, and the resulting influx of Cl ions hyperpolarizes the postsynaptic cell, thereby causing inhibition. During the past two decades, following affinity purification from mammalian spinal cord, the GlyRs have revealed much of their molecular organization, pharmacology, ligand-binding sites, associated disorders, and even three-dimensional structure, all of which are reviewed here.

2

Overview of GlyR Structure: Subunit Composition, Ligand-Binding Sites, and Ion Channel Region

2.1 Genes, Proteins, and Subunit Topology GlyRs are group I ligand-gated ion channels that belong to the Cys-loop receptor family, which also comprises the gamma-aminobutyric acid type-A (GABAA), 5-HT3, and nicotinic acetylcholine receptors (nAChR) (Lynch, 2004; Betz and Laube, 2006). Thanks to strychnine, a selective antagonist extracted from the Indian tree Strychnos nux vomica, the native GlyR was the first neurotransmitter protein to be isolated from the mammalian CNS by affinity purification (Pfeiffer et al., 1982). When analyzed by SDS-PAGE, the isolate was found to contain three distinct proteins: GlyR a (48 kDa) and b (58 kDa) subunits (Pfeiffer et al.,

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20

1982) in addition to a cytosolic scaffolding protein (93 kDa) named gephyrin (Prior et al., 1992). Crosslinking experiments demonstrated that native GlyRs are pentameric proteins, similar to the nAChRs isolated from the Torpedo electric ray (Langosch et al., 1988). Oligonucleotides deduced from the peptide sequences of purified subunits and homology screening were used to identify four different vertebrate genes (called Glra1–Glra4) encoding GlyR a subunits (a1–a4) and a single gene encoding the GlyR b subunit (Glrb) (Grenningloh et al., 1987, 1990a,b; Kuhse et al., 1990b; Matzenbach et al., 1994). All a subunits display high sequence identity (80–95%) and, upon heterologous expression, form functional homomeric glycine-gated channels with properties closely resembling those of native GlyRs. Biochemically, a subunits were shown to possess the critical determinants for ligand binding using photoaffinity labeling of the GlyR with [3H]-strychnine (Graham et al., 1983). The b subunit shows significant sequence differences as compared with the a subunits ( Figure 20-1) and might help in the rational design of novel selective agents. Classical agonists and competitive antagonists are known to bind to the ECD at the interface of two adjacent subunits (a–a for homomeric receptors; ab or ba for heteromeric receptors) (Betz and Laube, 2006). The pharmacology of the b–b interface is not known, but from studies using chimeric mutants involving the ECD of the b subunit and the transmembrane domain of the a1 subunit, fully functional homopentameric receptors were obtained, with properties comparable with wild-type a1 GlyRs, implying a functional b–b subunit interface (Kuhse et al., 1993; Griffon et al., 1999). The GlyR binding pocket is made of distinct ‘‘loop regions’’ (comprising random coil and b-strand structures) from the principal (+) and complementary () components (> Figure 20-1). Identified residues that affect agonist and antagonist binding include a1F63/R65 (loop D), a1E157/F159/G160/Y161 (loop B), a1R119 (loop E), and a1K200/Y202/T204/F207 (loop C) (Kuhse et al., 1990a; Vandenberg et al., 1992a, b; Schmieden et al., 1993, 1999) (> Figure 20-1). In addition, a disulfide bond between a1C198 and a1C209 constraining loop C appears critical for cell surface expression but also for ligand binding, most likely due to an indirect structural role (Rajendra et al., 1995b). Comparatively, the binding determinants of the b subunit have been little studied, despite a likely role in ligand binding (Rees et al., 2002). Only recently, two b subunit residues (R86 and E180, equivalent to a1R65 and a1E157, respectively) were shown to modestly (less than fivefold) influence ligand binding to heteromeric a1b GlyRs (Grudzinska et al., 2005). Docking of the agonist glycine and the antagonist strychnine into the pocket of GlyR ECD homology models resulted in complex structures that agreed well with the wealth of previous mutational data (Grudzinska et al., 2005) (> Figure 20-1).

2.3 Structure of the GlyR Transmembrane Domain The binding of agonists between the ECDs of GlyR subunits gates the opening of an anion-selective channel located within the transmembrane domain,  50A˚ away from the ECD. This transmembrane domain is composed of four tightly packed hydrophobic a helices arranged in a clockwise order (> Figure 20-1).

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. Figure 20-1 GlyR structure models and sequence relationships. (A) Homology model of the homopentameric a1 glycine receptor (side view). (B) Close-up view of the agonist binding site of the a1b GlyR with glycine molecule bound. (C) As B but bound to the antagonist strychnine. (D) Top view of the pentameric GlyRa1 ECD; histidine residues contributing to the inhibitory Zn2+ binding sites are highlighted in red. (E) Side view of the pore-forming TM2 segments of the receptor. Contacts between hydrophobic side chains of the TM2 segments at positions ‘9/’10 and ‘13/’14 (for details, see Miyazawa et al., 2003) are highlighted in red and yellow. The proline residues located at position 2 preceding the cytoplasmic end of TM2 are also depicted. (F) Phylogram of mammalian GlyR proteins. Phylogenetic scores show evolutionary distances between the receptor subunits. Panels B and C are reproduced with permission from Grudzinska et al. (2005)

TM1 to TM2, and TM2 to TM3, are connected by short loops, while TM3 to TM4 are linked by a long intracellular domain of about 100 residues, which includes phosphorylation and ubiquitination sites as well as a binding site for the scaffolding protein gephyrin (Meyer et al., 1995; Kneussel et al., 1999; Sola et al., 2004; Kim et al., 2006). While TM1, TM3, and TM4 isolate the pore region from the bilayer, stabilize the receptor in the membrane, and form a membrane-embedded binding site for alcohols and hydrophobic compounds (Cascio, 2006), several residues within the amphiphilic TM2 segment have been shown to line the ion channel. For easy comparison between different members of the Cys loop receptor family, the residues of TM2 are often numbered from 10 to 190 , with residue 10 corresponding to the cytoplasmic N-terminal end, and residue 190 to the extracellular end, of TM2 (Miller, 1989). When probed with the substituted cysteine accessibility method (SCAM), residues G20 , T60 , and R190 were labeled, consistent with these residues being pore lining in the a1-GlyR (Lynch et al., 2001). Additional pore-lining

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residues were inferred by sequence comparison with other LGICs and include T70 , L90 , T100 , T130 , S160 , and G170 (Xu and Akabas, 1996). For the nAChR, it has been suggested that TM2 is kinked inwardly at position L90 , which thereby should restrict the channel lumen (Miyazawa et al., 2003). However, the open-channel blocker QX-222, when applied in the open state of the channel, labeled also position 60 , implying that the narrowest portion of the channel is located relatively close to the cytoplasmic surface of the lipid bilayer (Charnet et al., 1990). Whether this is also true for the GlyR is unclear, but the general architecture of the Cys-loop receptor channel as derived from nAChR cryoelectron microscopy (Miyazawa et al., 2003) predicts that all TMs, and in particular TM4, extend significantly into the extracellular space, thus making them accessible for interaction with the ECD. The channel gate is thought to be located either in the central part of the pore, associated with a kink in TM2 (Miyazawa et al., 2003), or in the –20 –20 region, which forms the narrowest region of the pore according to SCAM studies (Xu and Akabas, 1996) and has been associated with the putative selectivity filter (Wilson and Karlin, 1998). Several residues outside of these regions have been shown to participate in the gating process, such as the GlyRa1 residues R271, located in the M2–M3 linker (Rajendra et al., 1995a), and K276, a pore-facing residue in the extracellular end of TM2 (Lewis et al., 1998). The mechanism of and the structural changes involved in channel gating are still unclear, but may include rotation of TM2 around position 90 (Unwin, 2005).

2.4 Ion Selectivity and Drug-Binding Sites Within the Transmembrane Region The glycine receptor channel is principally anion selective, but has been reported to also allow some permeation of sodium and potassium ions (Keramidas et al., 2002). In cultured spinal neurons, GlyRs display a permeability sequence of SCN>NO3>I>Br>Cl (Fatima-Shad and Barry, 1993), which is inversely correlated to the hydration energies of these ions, implying that removal of hydration water molecules is required for ion permeation (Bormann et al., 1987). Electrostatic potentials within the channel region are thought to contribute substantially to ion selectivity. The intracellular ends of the TM2 segments lining the pore of the GlyR provide a ‘‘ring of charges’’ consisting of arginine residues (R00 ), which are thought to concentrate permeating chloride ions and to repel oppositely charged cations. Consistent with this view, the subunits of cation-conducing nAChRs have a negatively charged residue – E10 at the equivalent position. In agreement with ion selectivity being regulated by electrostatic forces, a single point mutation (A-10 E) was found to suffice for converting the anion-conducing GlyR into a cationselective receptor (Keramidas et al., 2002). Additional rings of charge are provided by residues located at the extracellular ends of the TM2 segments and in the adjacent loops (R190 and K240 ), which upon mutagenesis were shown to reduce the unitary Cl conductance (Langosch et al., 1994). Finally, the a subunits of the GlyR contain a proline residue at position 20 , which has been suggested to play an important role in determining the selectivity and functional properties of the channel because of spatial disruptions around the pore boundaries (Gunthorpe and Lummis, 2001). While deletion of P20 alone does not alter ion selectivity, it was found to increase the pore size (from 5.4A˚ for wild-type to 6.9A˚ for the mutant channel) and to reduce the anion/cation permeability ratio (from 27 to 4) (Lee et al., 2003). In addition, a double substitution (the –P20 deletion combined with the A10 E mutation) significantly enhanced the relative cation permeability of the GlyR as compared with the single A-E10 mutant, confirming that in addition to electrostatic effects the pore size contributes to ion charge selectivity (Keramidas et al., 2002). Recently, dynamics simulations have been applied to GlyR-derived models of the nAChR ion pore structure; this revealed an energetic barrier of 7 kcal/mol in the center of the channel for sodium ions, whereas chloride ions were predicted to pass easily (Ivanov et al., 2007). Various compounds are thought to interact with the transmembrane domain of the GlyR (Lu and Xu, 2002; Cascio, 2006). Some of these, including channel blockers and compounds of the picrotoxin family, were shown by site-directed mutagenesis to bind to pore-exposed residues of TM2, though, in case of picrotoxin, mutation of several different and fairly distant residues reduced the efficacy of this compound, consistent with the existence of several distinct binding sites (Pribilla et al., 1992; Lynch et al., 1995; Dibas et al., 2002). In a recent study (Yang et al., 2007), different electrophysiological methods were used to

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20

demonstrate that positions 20 and 60 of TM2 provide important picrotoxin interaction sites. Another binding site for hydrophobic compounds, volatile anesthetics, and alcohols that potentiate GlyR function (Laube et al., 2002) is formed by domains TM1, TM3, and TM4. Due to its predicted size and shape, this site has been called ‘‘big cavity.’’ The mechanism of GlyR modulation by big-cavity binding compounds is not understood, despite its pharmacological importance. Mutagenesis and modeling approaches should help to clarify the nature and functional role of this intramembrane binding pocket.

3

Expression and Functions of GlyR Subtypes

3.1 GlyR Genes Display Distinct Expression Patterns in the Mammalian CNS Biochemical and in situ hybridization studies have shown that GlyR a1 mRNA and protein are prominently expressed in spinal cord, brain stem, and colliculi of adult rodents, whereas GlyR a2 protein and transcripts are abundant at birth and found only at low levels in adult hippocampus, cerebral cortex, and thalamus (Becker et al., 1988; Malosio et al., 1991; Sato et al., 1992). Moderate levels of GlyR a3 transcripts are detected in spinal cord, cerebellum, and olfactory bulb (Malosio et al., 1991). Due to its low abundance, GlyR a4 mRNA has so far not been localized in the mammalian CNS (Harvey et al., 2000). However, synaptic a4 subunit immunoreactivity has recently been visualized in the retina, a region of the CNS that expresses all known GlyR subunits (Heinze et al., 2007). The GlyR b (Glrb) gene is broadly expressed throughout both the embryonic and postnatal CNS (Malosio et al., 1991). In contrast, the GlyR a (Glra) genes show distinct developmental regulation: a1 GlyRs accumulate, and a2 GlyRs decrease, after birth (Becker et al., 1988; Malosio et al., 1991). Expression of a3 GlyRs also occurs postnatally (Malosio et al., 1991), whereas in the avian CNS a4, like a2, transcripts are mainly detectable at embryonic stages (Harvey et al., 2000). In conclusion, the expression of the Glra1–Glra4 genes is developmentally and regionally regulated in the mammalian CNS.

3.2 GlyRs Are Excitatory During Early Development Although glycine causes hyperpolarization, and thereby inhibition, of adult neurons, it serves as an excitatory transmitter during embryonic development and around birth. This is due to a positive chloride equilibrium potential of the developing neurons; therefore, GlyR activation results in chloride efflux causing depolarization (Reichling et al., 1994). This excitatory function of GlyRs at early developmental stages is important for synaptogenesis, since the GlyR-triggered activation of voltage-gated Ca2+ channels has been found to be crucial for proper formation of postsynaptic GlyR clusters (Kirsch and Betz, 1998). Postnatally, the chloride equilibrium potential shifts to negative values, because of active chloride extrusion by the K+/Cl cotransporter KCC2 (Rivera et al., 1999). Thereby GlyR currents become hyperpolarizing, that is, inhibitory. The developmental switch from excitatory to inhibitory GlyR function is paralleled by changes in receptor subunit composition. Embryonic and neonatal GlyRs are thought to be homopentamers of a2 subunits that are localized extrasynaptically (Hoch et al., 1989; Takahashi, 2005), whereas adult synaptic GlyRs are composed of a1 (or other a) and b subunits (Langosch et al., 1988; Meyer et al., 1995). The change in subunit composition results in characteristic alterations of the biophysical properties of postsynaptic GlyR currents, such as a smaller channel conductance and faster decay kinetics (Takahashi et al., 1992; Bormann et al., 1993; Singer et al., 1998). Recombinant homo-oligomeric a2 GlyRs show slower response kinetics and larger subconductance state distributions than a1b heteromeric receptors, consistent with the in vivo properties of neonatal versus adult GlyRs (overview in Takahashi, 2005). In other words, the transition from neonatal homo-oligomeric a2 GlyRs to adult hetero-oligomeric ab receptors underlies the shortening of the mean channel open time and the faster decay of inhibitory glycinergic postsynaptic currents seen at early postnatal stages.

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In addition to glycine, the amino acid taurine is an endogenous GlyR agonist. Taurine is released nonsynaptically from immature cortical neurons and has been reported to influence cortical development by activating excitatory extrasynaptic GlyRs (Flint et al., 1998). Furthermore, in in vitro preparations of the developing retina, the number of rod photoreceptors has been found to be regulated by taurine via a2 subunit-containing GlyRs expressed on retinal progenitor cells (Young and Cepko, 2004). Based on these data, extrasynaptic a2 GlyRs have been implicated in the regulation of neuronal development. However, recent studies on GlyR a2-deficient mice have failed to reveal deficits in brain anatomy, cortical development, and photoreceptor numbers (Young-Pearse et al., 2006; Weiss et al., 2008). Similarly, mouse lines lacking functional Glra1 and Glra3 genes show no alterations in CNS development and anatomy (Buckwalter et al., 1994; Harvey et al., 2004). These genetic data argue against an essential role of GlyRs in neuronal differentiation.

3.3 Roles of GlyR Isoforms in the Adult CNS: Lessons from Mutant Mice Due to the lack of subtype-specific antagonists, the physiological functions of the different GlyR isoforms have mainly been deduced from the analysis of GlyR-deficient mice. This has led to the following picture of GlyR subtypes.

3.3.1 a1 GlyRs GlyRs containing the adult a1 subunit represent the major GlyR isoform. a1 GlyRs are expressed in many regions of the CNS, and their blockade explains the major physiological symptoms of strychnine poisoning, such as convulsions due to disinhibition of motor neurons and hyperexcitability resulting from disinhibition of sensory afferents. Coassembled with the b subunit, a1b receptors account for the majority of synaptically localized GlyRs in the mammalian CNS, and hence are enriched in receptor preparations purified by strychnine affinity chromatography (Pfeiffer et al., 1982; Grenningloh et al., 1987). The importance of the GlyR a1 subunit is further underlined by the existence of disease mutations in the Glra1 genes of different mammalian species, including mouse, cattle, and human (Lynch, 2004; see below). The spasmodic mouse carries a point mutation at position 52 (A52S) of Glra1 that reduces the agonist binding affinity (Ryan et al., 1994; Saul et al., 1994). The oscillator strain constitutes a natural Glra1 knockout mouse, because of a microdeletion in exon 8 (Buckwalter et al., 1994). Although homozygous spasmodic mice survive into adulthood with persistent symptoms of spasticity, myoclonus, and hyperexcitability, oscillator homozygotes die at the end of the third postnatal week, because of clonic convulsions and extreme muscle stiffness. Upon recombinant expression, the GyR a1 subunit forms channels characterized by short mean open times and fast decay kinetics, as found for glycinergic spontaneous inhibitory postsynaptic currents (sIPSCs) in adult spinal cord neurons (Singer et al., 1998). Moreover, in retina preparations obtained from oscillator mice glycinergic sIPSCs in A-type ganglion cells are strikingly reduced, and their kinetics slowed, because of Glra1 inactivation (Majumdar et al., 2007). Together, these findings indicate that the GlyR a1 subunit defines receptors that are crucial for fast regulation of both motor and sensory functions.

3.3.2 a2 GlyRs Despite the recent availability of Glra2-deficient mice (Young-Pearse et al., 2006; Weiss et al., 2008), the precise physiological roles of a2 GlyRs are still enigmatic. At early developmental stages, the GlyR a2 subunit is known to form homo-oligomeric receptors (Hoch et al., 1989), which are extrasynaptically localized and may mediate non-synaptic tonic transmission caused by non-vesicular glycine release and/or spillover from adjacent nerve terminals. Consistent with the role of a2 GlyRs as ‘‘tonic receptors,’’ a major fraction of the

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20

a2 immunoreactivity found in hippocampal neurons has been shown to cluster at sites that do not colocalize with other marker proteins of inhibitory synapses (Danglot et al., 2004; Papadopoulos et al., 2007). Synaptically localized a2 staining has been detected in different neuronal cell populations including amacrine cells of the retina and is thought to correspond to a2b heteromeric receptors (Haverkamp et al., 2004). Electrophysiological studies of neonatal GlyRs in spinal neurons (Singer et al., 1998) and, recently, glycinergic sIPSCs in narrow field amacrine cells of Glra2-deficient mice (Weiss et al., 2007) strongly support the idea that GlyRs containing the a2 subunit mediate inhibitory postsynaptic currents characterized by slow decay kinetics.

3.3.3 a3 GlyRs Originally, the a3 subunit was considered a minor GlyR protein with an expression pattern similar to that of the a1 subunit (Kuhse et al., 1990b; Malosio et al., 1991). More recently, immunocytochemistry with specific antibodies revealed that in spinal cord a3 GlyRs are selectively localized at synaptic sites in the superficial laminae I and II of the dorsal horn, where nociceptive afferents terminate (Harvey et al., 2004). Here, a local network of glycinergic interneurons inhibits the propagation of nociceptive signals to higher brain regions. The analysis of knockout mice has shown that the a3 GlyRs in the dorsal horn represent the molecular substrate of pain sensitization by the inflammatory mediator prostaglandin E2 (PGE2). PGE2 activates prostaglandin E receptors of the EP2 subtype and leads to a protein kinase A-dependent downregulation of glycine currents in dorsal horn neurons (Ahmadi et al., 2002). Upon inactivation of the murine Glra3 gene, this down-regulation of glycine inhibition by PGE2 is abolished, and PGE2 fails to sensitize pain responses in Glra3 knockout animals (Harvey et al., 2004). Therefore, a3 GlyRs are considered a promising target for designing novel therapeutic strategies to achieve analgesia, for example, by administering drugs that selectively potentiate a3 currents or by enhancing the RNA editing process that generates high-affinity a3 GlyRs (Meier et al., 2005). GlyR a3 transcripts have also been detected in CNS regions other than spinal cord (Malosio et al., 1991). In retina, GlyRa3 is found at synapses that are largely distinct from GlyR a1-containing ones (Haverkamp et al., 2003). By analyzing GlyR a3-deficient mouse lines, synaptic GlyRs of AII amacrine cells have recently been shown to contain a3 subunits that confer medium–fast kinetics on sIPSCs (Weiss et al., 2008). Together, the presently available data are consistent with a3 GlyRs being preferentially expressed in sensory pathways.

3.3.4 a4 GlyRs The a4 GlyRs are the least understood members of the GlyR family. Although discovered more than a decade ago by genomic screening (Matzenbach et al., 1994), it still is unknown whether and where the Glra4 gene is expressed, and what its functions are in the mammalian CNS. This mainly reflects the low abundancy of a4 mRNA and protein. In the chicken, a4 transcripts have been detected at embryonic stages in spinal cord, peripheral ganglia, and the male genital ridge (Harvey et al., 2000). Very recently, a4 immunoreactive synapses have been visualized on displaced ON cholinergic amacrine cells in the adult rodent retina (Heinze et al., 2007). Upon heterologous expression, a4 GlyRs form channels with pharmacological properties largely resembling those of a1 receptors (Harvey et al., 2000). Notably, in humans the Glra4 gene may be a pseudogene. Based on the analysis of sIPSCs in narrow field amacrine cells of GlyR a2-deficient mice, a4 receptors have been speculated to generate channels displaying ultra-slow decay kinetics (Weiss et al., 2007).

3.3.5 GlyR b Subunit As outlined earlier, the ubiquitously expressed b subunit is essential for the formation of synaptic GlyRs, since it provides for receptor anchoring to the synaptic scaffolding protein gephyrin (Kneussel and Betz,

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2000). Its importance is also underlined by the spastic mutation in mice, which produces a phenotype identical to that seen in spasmodic animals (White and Heller, 1982). Spastic animals have normal synaptic a1b GlyrRs but an intronic insertion of a LINE element in the Glrb gene results in a strong reduction of receptor expression and aberrant splicing of the b pre-mRNA (Becker et al., 1986, Kingsmore et al., 1994; Mu¨hlhasdt et al., 1994). Glrb knockout mice have not been produced but may display perinatal lethality due to a lack of synaptic receptors, as found in gephyrin-deficient animals (Feng et al., 1998).

4

Human Disorders of Glycine Neurotransmission

4.1 Mutations in GlyR Genes Result in Hereditary Neuromotor Disease The first disorder associated with a mutation in a GlyR gene was found by positional cloning. Hyperekplexia (startle disease, Kok disease, OMIM149400) is characterized by muscle rigidity of nervous system origin, particularly in neonates, and by an exaggerated startle response to unexpected auditory or tactile stimuli. Consciousness, however, remains intact. Clonazepam, a positive modulator of GABAA receptormediated inhibitory transmission, provides symptomatic relief. In 1993, the underlying genetic defect was mapped to chromosome 5q33-q35 by genetic linkage using DNA markers defining the GLRA1 gene as the prime candidate (Shiang et al., 1993). Mutations resulting in the substitution of the highly conserved residue R271 by uncharged amino acids (L or Q) were identified in four families with autosomal dominant hyperekplexia. These substitutions result in a reduction of agonist sensitivity and glycine-activated current. Recombinant expression of the mutant GlyRs revealed that the reduced glycine current is due to a decrease in single-channel conductance levels. An in vivo study using transgenic mice expressing the human GlyR a1R271Q disease protein revealed diminished glycinergic neurotransmission, resulting in exaggerated startle responses and myoclonus (Becker et al., 2002), which could be transiently cured by subanesthetic concentrations of propofol, an intravenous anesthetic that potentiates GlyR currents (O’Shea et al., 2004). Subsequently, additional autosomal dominant and recessive inheritance patterns and compound heterozygosity have also been described in other patient families that carry missense or null mutations in the GLRA1 locus (see > Table 20-1; Rees et al., 1994; Brune et al., 1996; Humeny et al., 2002; Gilbert et al., 2004; Tsai et al., 2004; Coto et al., 2005). However, in less than half of the >70 pedigrees of human patients displaying hyperekplexia-like symptoms investigated presently, GLRA1 mutations have been identified. This suggests that genes other than GLRA1 could be involved in hyperekplexia. Consistent with an important role of the b subunit and the scaffolding protein gephyrin in postsynaptic GlyR function, mutations in the GLRB (Rees et al., 2002) and GPHN (Rees et al., 2003) genes have been found to be associated with hyperekplexia > Table 20-1. In addition, the gene encoding the neuronal glycine transporter GlyT2 (SLC6A5) has been recently shown to be a major site of hyperekplexia mutations (Eulenburg et al., 2006; Rees et al., 2006). So far, no pathogenetic mutations have been detected in the GLRA2 and GLRA3 genes. Recent studies have associated GlyRs with the pathology of autism (Ramanathan et al., 2004), HIVassociated dementia (Gelman et al., 2004), generalized epilepsy (Sobetzko et al., 2001), and amyotrophic lateral sclerosis (Lorenzo et al., 2006). For example, besides its role in inflammatory pain sensitization established by the analysis of GlyRa3 knockout mice (Harvey et al., 2004), the GLRA3 gene has been implicated in autism and idiopathic generalized epilepsies, based on polymorphisms and an interstitial deletion on chromosome 4q within the region encoding the GlyR a3 subunit (Sobetzko et al., 2001; Ramanathan et al., 2004). Future studies may thus unravel important roles of GlyR mutations in diverse neurological disorders besides their well-documented importance in hyperekplexia.

5

Conclusion and Perspectives

During the past decades, significant progress has been made in understanding the structures and specific physiological functions of distinct GlyR subtypes. Furthermore, mouse models for different GlyR subunit deficiencies have become available, and the importance of GlyR mutations in the pathogenesis of human

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. Table 20-1 Mutations associated with hyperekplexia Demonstrated functional effects Impaired membrane insertion Reduced agonist affinity and channel gating Reduced channel function

Mode of inheritance D

Gene GLRA1

Mutation R218Q

GLRA1

P250T

GLRA1 GLRA1

V260M Q266H

GLRA1 GLRA1

S270T R271Q,L

GLRA1

K276E

D

GLRA1

Y279C,S

D

GLRA1

G342S

D

GLRA1

S231R

GLRA1 GLRA1 GLRA1 GLRA1 GLRA1 GLRA1 GLRA1

R100H Y202X Y228C I244N deletion W96C/R344X R252H/R392H

GLRB GPHN SLC6A5

G229D/exon5 N10Y multiple

Reduced agonist affinity and channel conductance

Impaired membrane insertion Reduced expression Reduced protein level GlyR assembly Reduced protein level

Reduced agonist affinity

D D D D D

R R R R R R C C C D R

References Castaldo et al. (2004) Breitinger et al. (2001), Saul et al. (1999) del Giudice et al. (2001) Castaldo et al. (2004), Milani et al. (1996), Moorhouse et al. (1999) Lapunzina et al. (2003) Kimura et al. (2006), Langosch et al. (1994), Shiang et al. (1993) Elmslie et al. (1996), Lewis et al. (1998), Seri et al. (1997) Kwok et al. (2001), Poon et al. (2006), Shiang et al. (1995) Jungbluth et al. (2000), Rees et al. (2001) Humeny et al. (2002) Coto et al. (2005) Rees et al. (2001) Forsyth et al. (2007) Rees et al. (1994) Brune et al. (1996) Tsai et al. (2004) Rea et al. (2002), Vergouwe et al. (1999) Rees et al. (2002) Rees et al. (2003) Eulenburg et al. (2006), Rees et al. (2006)

D, autosomal dominant; R, autosomal recessive; C, compound heterozygosity

diseases is now clearly demonstrated. However, many open questions await further investigation, including the mechanism of GlyR channel gating, the precise localization of anesthetic binding sites, and the individual roles of the different GlyR subunits in various normal and diseased brain regions.

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Aquaporins in the Brain

M. Yasui . Y. Fujiyoshi

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 392

2 2.1 2.2 2.3 2.4 2.5

Expression and Distribution of AQP4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393 Distribution of AQP4 in Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393 Polarized Expression of AQP4 in Astrocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393 AQP4 Expression Outside Astrocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394 Ontogenic Expression of AQP4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 395 Altered Expression of AQP4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 395

3 3.1 3.2 3.3

Structures of AQP4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396 Common Structures of Aquaporins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396 Crystal Structure of AQP4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396 Square Array Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396

4 Regulation of AQP4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 398 4.1 Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 398 4.2 Inhibition of Metals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399 5 AQP4 KO Mice Phenotype . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399 5.1 Brain Edema . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399 5.2 Impaired Neuroexcitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400 6 AQP4 and Human Clinical Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 401 6.1 Neuromyelitis Optica (NMO) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 401 7

#

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 401

2009 Springer ScienceþBusiness Media, LLC.

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Aquaporins in the brain

Abstract: Discovery of aquaporin water channel proteins has provided insight into the molecular mechanism of membrane water permeability. In mammalian brain, Aquaporin-4 (AQP4) is the main water channel and is distributed with highest density in the perivascular and subpial astrocyte end-feet. AQP4 is a critical component of an integrated water and potassium homeostasis. Indeed, AQP4 has been implicated in several neurologic conditions, such as brain edema and seizure by phenotype analysis of AQP4-null mice. Expression and regulation of AQP4 have been studied to understand the roles of AQP4 in physiological and pathological conditions. AQP4 is dynamically regulated at different levels; channel gating, subcellular distribution, phosphorylation, protein-protein interactions and orthogonal array formation. Interestingly, AQP4 has been identified as a target antigen of autoimmune attack in neuromyelitis optica (NMO). AQP4 may be a potential therapeutic target in several neurologic conditions. Further studies from different aspects are required to develop new drugs against AQP4. List of Abbreviations: AQP, aquaporin; CSF, cerebrospinal fluid; CKII, casein kinase II; CaMKII, calcium/ calmodulin dependent protein kinase II; DG, dystroglycan; ECS, extracellular space; FRL, freeze fracture replica labeling; PDZ, PSD95-Discs large-ZO1; PKA, protein kinase A; PKC, protein kinase C; NMO, neuromyelitis optica; Syn, syntrophin

1

Introduction

Water constitutes approximately 70% of our body mass; an appropriate distribution of water is necessary to maintain an orderly fluid balance within different anatomic compartments. Therefore the regulated movement of water across cell membranes is fundamental for life. While the water permeability of most cell membranes can be accounted for by the flip-flop motion of lipid molecules, certain cells (i.e., red blood cells and kidney proximal tubule cells) exhibit membrane water permeability far too great to be explained solely by such a mechanism (Finkelstein, 1987). These observations have led to speculation about the existence of water-specific channels or pores in some tissues. The first water channel, aquaporin-1 (AQP1), was discovered by Agre’s group during studies of human red blood cell Rh proteins. Cloning of the full-length cDNA revealed significant homology with the major intrinsic protein (MIP) of the lens (Preston and Agre, 1991). Expression of AQP1 in Xenopus laevis oocyte provided evidence that AQP1 is a water-specific channel as shown in > Figure 21-1 (Preston et al., 1992). Since the discovery of AQP1, hundreds of homologous proteins have been recognized in plants, microbials, invertebrates, and vertebrates. Thirteen mammalian aquaporins have been identified to date, each with a distinct tissue distribution (Borgnia et al., 1999). Multiple regulatory mechanisms are likely to be identified that will explain the specificity of ontogeny, distribution, and function of each aquaporin (Agre et al., 2002). Water movement in the brain is not well understood although about 85% of it is water (Tait et al., 2008). Water transport is directly or indirectly coupled to a number of other homeostatic processes in the brain, such as potassium clearance and glutamate uptake. Brain water transport is important in pathological conditions such as brain edema and hydrocephalus. Among mammalian aquaporins, AQP1, AQP4, and AQP9 are expressed in the brain (Amiry-Moghaddam and Ottersen, 2003). AQP1, expressed in the apical membrane of the choroid plexus, may play a role in secreting cerebrospinal fluid (CSF). Indeed, AQP1 is upregulated in choroid plexus tumors, associated with increased CSF production (Longatti et al., 2006). On the other hand, AQP1 null mice did not reveal a significant decrease in CSF volume (Oshio et al., 2005). Expression and distribution of AQP9 in the brain is still controversial. AQP9 immunoreactivity in the rat or mouse brain has been observed in tanycytes (a subpopulation of glia) and some population of catecholaminergic neurons (Badaut et al., 2004). However, comparable AQP9 immunoreactivity was also observed in AQP9-null mice, suggesting that AQP9 antibodies might crossreact with other proteins (Rojek et al., 2007). In this chapter, we review AQP4, the predominant aquaporin in the brain.

Aquaporins in the brain

21

. Figure 21-1 Functional expression of AQP1 water channel in Xenopus oocytes. Incubation in hypotonic buffer leads to swelling of an oocyte injected with AQP1 cRNA, indicating high osmotic water permeability (right). In contrast, a control oocyte (left) fails to swell. Reprinted from Science with permission (Preston et al., 1992)

2

Expression and Distribution of AQP4

2.1 Distribution of AQP4 in Brain AQP4 is distributed throughout the brain and spinal cord as well as the cerebellum (Nielsen et al., 1997). AQP4 has not been found in neurons but has been exclusively found in the plasma membrane of astrocytes. AQP4 is also expressed in the basolateral membrane of ependymal cells lining ventricles. Interestingly, astrocytes exhibit highly polarized expression of AQP4, which is concentrated in astrocyte end-feet membranes that are in direct contact with blood vessels, the ependymal layer and pia. This pattern of distribution suggests that AQP4 may play a role in brain water homeostasis by controlling water flow into and out of the brain (> Figure 21-2).

2.2 Polarized Expression of AQP4 in Astrocytes The polarized expression of AQP4 in the astrocyte end-feet facing capillary endothelial cells is altered in the absence of endothelia, i.e., malignant astrocytes and cultured astrocytes, suggesting that endothelial cells signal astrocytes to polarize AQP4 expression (Saadoun et al., 2002). When astrocytes are co-cultured with endothelial cells, the two cell types directly contact each other, and AQP4 is redistributed in the astrocyte processes in close contact with the endothelial cells (Nicchia et al., 2004). In addition, the polarized expression of AQP4 requires a-syntrophin in the astrocyte since AQP4 expression is significantly reduced in astrocyte end-feet adjacent to blood vessels in the cerebral cortex of a-syntrophin-null mice (Neely et al., 2001; Amiry-Moghaddam et al., 2003b). AQP4 has a C-terminal PDZ (PSD95-Discs large-ZO1)-binding motif (Ser-Ser-Val). It has been

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. Figure 21-2 Schematic of AQP4 distribution and routs of water flow out of the brain. AQP4 expression at barriers in the exit routes of fluid throughout the brain and spinal cord as well as the cerebellum. Excess of fluid moves from brain: through several layers of astrocyte processes (the glia limitans externa) into the subarachnoid space (top left); through astrocyte foot processes and endothelial cells into the bloodstream (top right); through the glia limitans interna and ependyma into the ventricles (bottom left). Reprinted from FASEB J. with permission (Papadopoulos et al., 2004)

suggested that expression and subcellular localization of AQP4 depend on association with the dystrophin complex through a tSSV-PDZ-medicated interaction with a-syntrophin (> Figure 21-3). Mutations in the human gene encoding dystrophin are found in patients with Duchenne muscular dystrophy, and the mdx mouse is an animal model for this important disorder. AQP4 expression in muscles has been significantly reduced in these cases (Frigeri et al., 2001; Wakayama et al., 2002). On the other hand, the physiological relevance of AQP4 in muscle tissue is largely unknown, and more studies are needed.

2.3 AQP4 Expression Outside Astrocytes AQP4 is also expressed in the supraoptic nucleus and circumventricular organs, such as the area posterma and the vascular organ of the lamina terminals, which lack a blood-brain barrier and contain osmosensitive magnocellular neurons that regulate fluid homeostasis and release antidiuretic hormone (Nielsen et al., 1997). In the cerebellum, AQP4 is expressed in glial cells; mainly in radial processes (Bergmann fibers) and the subpial and perivascular astrocyte end-feet (Wen et al., 1999). In the optic nerve, AQP4 is expressed in Muller cells in the retina and fibrous astrocytes (Nagelhus et al., 1998).

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. Figure 21-3 AQP4 in the dystrophin-associated protein complex. The dystrophin-associated protein complex is a large membrane assembly that connects the cytoskeleton to the extracellular matrix. AQP4 is polarized at this site through direct or indirect interaction with other proteins through its C-terminal PDZ-binding motif (Ser-Ser-Val). Reprinted from Nature Reviews (Neuroscience) with permission (Amiry-Moghaddam and Ottersen, 2003)

2.4 Ontogenic Expression of AQP4 As for ontogeny, AQP4 expression is limited in the first week of postnatal life, increases significantly during the second and third weeks, and then reaches adult levels after 4 weeks (Wen et al., 1999). In addition, AQP4 is expressed during the blood-brain barrier formation in chick embryos (Nicchia et al., 2004).

2.5 Altered Expression of AQP4 Altered expression of AQP4 has been observed both in vivo and in vitro models, suggesting some physiological and pathological relevance for AQP4. In general, up-regulation of AQP4 expression is associated with brain injuries and tumor formation. AQP4 is highly expressed in human gliomas and is associated with blood-brain barrier disturbance (Warth et al., 2007). Biphasic changes in AQP4 expression were observed in spinal cord injury rat models: early down-regulation and subsequent persistent upregulation (Nesic et al., 2006). There is a report showing the up-regulation of AQP4 by intracerebral VEGF

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injection, which has beneficial effects in ischemic rats (Rite et al., 2008). AQP4 is also expressed in primary astrocyte cell cultures, although polarized distribution is no longer maintained. AQP4 expression in primary astrocytes is up-regulated in response to various stimuli, including P2X7 receptor activation, hypertonic shock, and agrin, heparin sulfate proteoglycan of the extracellular matrix (Lee et al., 2008). Further studies are required to understand the molecular mechanisms behind the up-regulation of AQP4 expression.

3

Structures of AQP4

3.1 Common Structures of Aquaporins The deduced amino acid sequence of AQP confirmed that protein most likely constitutes six bilayerspanning domains with intracellular N- and C-terminal domains. Based on biochemical and site-directed mutagenesis studies, an ‘‘hourglass model’’ was proposed: a pseudo two-fold symmetrical structure with six bilayer-spanning a-helices surrounding the aqueous pore formed the two NPA (Asn, Pro, Ala)-containing loops that enter the bilayer from the opposite surfaces and overlap at the contact point of the two NPA motifs (Jung et al., 1994). Several years later, it was shown that this ‘‘hourglass model’’ is basically correct by demonstrating the high-resolution atomic model of AQP1. The structure of human AQP1 reconstituted into membrane crystals was determined at 3.8 A˚ by cryoelectron microscopy; the structure of three-dimensional bovine AQP1 was obtained at 2.8 A˚ by X-ray diffraction analysis (Murata et al., 2000; Sui et al., 2001). These structures are remarkably uniform and explain how aquaporin functions in terms of selectivity and conductance at an atomic level (Kozono et al., 2002). All known aquaporin structures reveal a common homo-tetrameric arrangement with each monomer folded into a right-handed helical bundle architecture. Unlike the potassium channel, in which four subunits surround a central pore, each monomer contains its own water-conducting pore (Murata et al., 2000).

3.2 Crystal Structure of AQP4 AQP4 structure has been determined by electron crystallography of double-layered, two-dimensional crystals at 3.2 A˚ (Hiroaki et al., 2006). AQP4 forms a homo-tetramer with a typical AQP fold of each monomer. The water-conducting pore structure of AQP4 is basically identical to that of AQP1. Unlike other AQPs, AQP4 contains a short 310 helix in an extracellular loop. Furthermore, the short 310 helix mediates specific interactions between AQP4 molecules in adjoining membranes. This unique structure of AQP4 suggests an unexpected role in cell adhesion as shown in > Figure 21-4. Indeed, AQP4 transfected L cells, which have no endogenous adhesive properties, formed aggregated clusters (Hiroaki et al., 2006).

3.3 Square Array Formation AQP4 uniquely forms higher oligomeric structures, the so-called square arrays, prominent in the plasma membranes of brain astrocytes, glial lamellae of hypothalamus, stomach, renal collecting duct, muscles, and retina where AQP4 is expressed (Yang et al., 1996). This suggests that AQP4 is involved in squarearray formation. Direct anti-AQP4 immunogold labeling identified square arrays as microcrystalline assemblies of AQP4 (Rash et al., 1998). Furthermore, the absence of square arrays in brains from AQP4 null mice clearly indicated that the square arrays are formed by AQP4 (Verbavatz et al., 1997). AQP4 has two alternative splice variants resulting from differential translation initiating methionines, M1, or M23 isoform (Lu et al., 1996). M1 has an N terminus 22 residues longer than M23. It has been demonstrated that AQP4 forms heterotetramers by both isoforms. Freeze-fracture analyses indicate that M23 forms large square arrays while M1 does not form square arrays when transiently expressed in mammalian cells. When both M1 and M23 are co-expressed, they make the intermediate size of arrays, resembling

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. Figure 21-4 Adhesive structure of two AQP4 molecules in adjoining membranes. The interaction between AQP4 monomers in the adjoining membranes. The AQP4 molecules are represented by ribbon diagram; H1-H6, transmembrane helices; HB, HE, pore helices in loops B and E; HC, 310 helix in loop C. The residues of Pro139 and Val142 in the 310 helices, responsible for junction interactions, are shown in ball-and-stick representation

square arrays of in vivo astrocytes (Furman et al., 2003). These observations suggest that the M1 isoform restricts square-array assembly and that the additional 22 N-terminal residues of M1 interfere with array formation. The structure of AQP4M23, determined by electron crystallography, revealed the array-forming mechanism, such as the characteristic intermolecular interaction mediated by Arg and Tyr residues in the AQP4 2D-crystals, which resembled the one seen in the square arrays (Hiroaki et al., 2006). Site-directed mutagenesis, the SDS-digested freeze-fracture replica labeling technique, and metabolic labeling experiments also revealed that the two cysteine residues at positions 13 and 17 in the N terminus of M1 are palmitoylated and thereby interfere with array formation (> Figure 21-5) (Suzuki et al., 2008). Since there is no significant difference in water permeability between M1 and M23 when expressed in Xenopus oocytes, the biological significance of the array formation is not yet clear (Neely et al., 1999). So far, there is no evidence that the size of a square array is dynamically regulated in response to various stimuli.

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. Figure 21-5 SDS-FRL images of CHO cells transfected with four types of AQP4. Square array formation and deformation of mutants of AQP4 were observed by SDS-digested freeze fracture replica labeling (SDS-FRL). The C-terminal of AQP4 molecules are labeled by anti-AQP4CT antibody and immunogold particles indicate the AQP4 molecules in the SDS-FRL images. The SDS-FRL images of CHO cells transfected with AQP4M1, AQP4M23, AQP4M1/C13A and AQP4M1/C13,17A are represented in a, b, c, and d, respectively. From these data, tow Cys residues, C13 and C17 were confirmed to be important for interfering array formation of AQP4. Scale bars are 200 nm

4

Regulation of AQP4

4.1 Phosphorylation Several studies have demonstrated that AQP4 may be regulated by phosphorylation. AQP4 has several putative phosphorylation sites for protein kinase A (PKA), protein kinase C (PKC), calcium/calmodulindependent protein kinase II (CaMKII) and casein kinase II (CKII) (Gunnarson et al., 2004). Osmotic water permeability of AQP4 is dose-dependently inhibited by phorbol esters when expressed in Xenopus oocytes (Han et al., 1998). The water permeability of AQP4 is also downregulated by PKC activation and by dopamine in mammalian epithelial cells transiently transfected with AQP4 (Zelenina et al., 2002). Site-directed mutagenesis reveals that Ser180 is the site of PKC phosphorylation. However, it is still not clear how PKC phosphorylation leads to the inhibition of AQP4 activity – by channel gating or by trafficking. In contrast, the water permeability of astrocyte membranes expressing AQP4 is increased by lead via the CaMKII pathway (Gunnarson et al., 2005). There is a report that AQP4 internalization is followed by an increase in AQP4 phosphorylation in response to histamine through activation of PKA in AQP4-transfected gastric HGT1 cells (Carmosino et al., 2007). Phosphorylation of Ser276 by CKII enhances AQP4-lysosomal targeting and degradation (Madrid et al., 2001). Overall, regulation of AQP4 by phosphorylation seems to be very complicated. More studies are needed to evaluate the biological relevance of AQP4 phosphorylation in an in vivo model.

Aquaporins in the brain

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4.2 Inhibition of Metals AQP4 was considered to be a mercury-insensitive AQP although most AQPs, including AQP1, are inhibited by mercurials (Hasegawa et al., 1994). Site-directed mutagenesis of AQP1 revealed that Cys189 at the constriction of the pore is a binding site for Hg2+ (Preston et al., 1993). AQP4 does not have a cysteine residue at the position corresponding to Cys189 of AQP1, supporting the concept that AQP4 is a mercuryinsensitive AQP. However, AQP4 has been recently shown to be inhibited by mercury, when reconstituted into proteoliposomes, in a manner different from that of AQP1 (Yukutake et al., 2008). In case of AQP4, Cys178, which is located in intracellular loop D, is a target responding to Hg2+. Interestingly, other metals, including silver and gold, also inhibit AQP4 via C178. Since C178 is far from the pore constriction, it is considered that Hg2+ or other metals binding to C178 induce conformational changes of AQP4, leading to the inhibition of the channel.

5

AQP4 KO Mice Phenotype

Phenotype analysis of AQP4-null mice has provided compelling evidence for roles of AQP4 in cerebral water balance (Verkman et al., 2006).

5.1 Brain Edema Water accumulation is the hallmark of brain edema. It is associated with several brain pathological conditions such as stroke, brain tumor, hydrocephalus, and traumatic injury, and is also seen in systemic pathological conditions including hyponatremia, sepsis, meningitis, and childhood cerebral malaria (Papadopoulos and Verkman, 2008). Brain edema results in elevated intracranial pressure, potentially leading to brain ischemia, herniation, and death. The therapies to reduce brain swelling have not been significantly improved for more than 50 years, and it is particularly urgent to develop new ones. The pattern of AQP4 distribution provides indirect evidence for involvement of AQP4 in the development of brain edema (Papadopoulos and Verkman, 2007). This is supported by evidence that AQP4 expression is lost from perivascular end-feet in the post-ischaemic phase, which might be a mechanism to limit water influx (Nase et al., 2008). Direct evidence has come from experiments showing reduced brain edema formation with improved survival rate in AQP4-null mice compared to wild-type mice after water intoxication (one example of cytotoxic edema where the blood-brain barrier is intact) (> Figure 21-6) (Manley et al., 2000). Intraperitoneal water injection causes profound hyponatremia and subsequent death from brain edema and increased intracranial pressure. Mortality after hyponatremia is markedly reduced in AQP4-null mice, associated with reduced brain water accumulation and astrocyte foot-process swelling in contact with blood vessels. Reduced brain swelling and improved outcome is also seen in AQP4-null mice in a model of ischemic stroke due to transient middle cerebral artery occlusion and in a model of bacterial meningitis induced by intracisternal streptococcus injection (Papadopoulos and Verkman, 2005). Reduced brain swelling is also shown in a-syntrophin null mice as well as in dystrophin null (mdx) mice, which secondarily disrupted AQP4 expression and distribution (Vajda et al., 2002; Amiry-Moghaddam et al., 2003b). Taken together, these findings with transgenic mice suggest that AQP4 can be a therapeutic target for brain edema in humans. In contrast, AQP4 null mice had more brain swelling compared with wild-type mice after cortical freeze injury and brain tumor implantation (Papadopoulos et al., 2004). These conditions are considered to be vasogenic edema, where the blood-brain barrier is disrupted. In these cases, AQP4 may play a role in eliminating excess fluid from brain parenchyma. Although brain edema is classified into several types, such as cytotoxic edema and vasogenic edema, most clinical conditions consist of mixtures of different types of edema with different time courses. For example, early cerebral ischemia produces astrocyte swelling, but later on, the

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. Figure 21-6 Effect of water intoxication on survival in AQP4 +/+ and AQP4 / mice. (a) Mice received intraperitoneal injection of distilled water (20% body weight) and the percentage of surviving AQP4 +/+ and AQP4 / mice is shown for each time point. (b) Quantification of cerebral edema following water intoxication. Transmission electron micrograph showing edematous cerebral cortex at 30 min. The swollen astrocyte foot-process in brain from AQP4 +/+ (white arrows) and AQP4 / (black arrows). Modified from Nature Med. with permission (Manley et al., 2000)

blood-brain barrier is disrupted, resulting in vasogenic edema. It is therefore not so simple to predict whether and when AQP4 could be inhibited or activated to reduce brain swelling.

5.2 Impaired Neuroexcitation In addition to the involvement in brain edema, AQP4 is implicated in pathological conditions associated with perturbed ion homeostasis. Several lines of evidence suggest that AQP4 controls the brain extracellular space (ECS) volume and K+ uptake by glial cells (Amiry-Moghaddam et al., 2004). AQP4 has been suggested to be coupled functionally and spatially to the potassium channel, Kir4.1, that is responsible for K+ buffering after neuronal activity (Nagelhus et al., 1999). Moving in parallel with K+, water must be siphoned into blood or CSF during periods of high neuronal activity. AQP4 may play a role in transporting water through astrocyte cell membranes. Verkman, et al. developed a method to measure ECS in vivo using fluorescently labeled macromolecules with surface photobleaching (Binder et al., 2004a, Zador et al., 2008). They found that ECS volume was increased in AQP4-null mice. Impaired K+ reuptake in AQP4 deficiency was observed using a long-wavelength K+-sensing fluorescent indicator, TAC-Red (Padmawar et al., 2005). Altered ECS volume and/or K+ handling resulted in impaired neural signal transduction in AQP4 deficiency, which is shown in several transgenic mouse models. (1) Seizure susceptibility in response to the convulsant pentylenetetrazol was reduced in AQP4-null mice (Binder et al., 2004b). (2) Electrically induced seizures following hippocampal stimulation showed greater threshold and remarkably longer duration in AQP4-null mice (Binder et al., 2006). (3) a-syntrophin-null mice developed more severe behavioral seizures after hyperthermia (Amiry-Moghaddam et al., 2003a). It is still unclear, however, how ECS volume is increased and K+ clearance is delayed, and if these two conditions are closely related to each other in AQP4-null mice. In addition, AQP4-null mice revealed increased auditory brainstemevoked response thresholds and reduced electroretinographic potential (Takumi et al., 1998; Li and Verkman, 2001).

Aquaporins in the brain

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AQP4 and Human Clinical Disorders

6.1 Neuromyelitis Optica (NMO) Neuromyelitis optica (NMO) is an inflammatory demyelinating disorder that predominantly affects the optic nerve and spinal cord (Wingerchuk et al., 2007). Recent serological findings strongly suggest that NMO is a distinct disease rather than a subtype of multiple sclerosis (MS), since Lennon et al. surprisingly demonstrated that a serum immunoglobulin G autoantibody (NMO-IgG) that selectively binds to AQP4 is a specific marker of this disorder (Lennon et al., 2005). NMO-IgG is detectable in the majority of patients with NMO (68–91%) and highly specific for NMO (up to 98%) (Lennon et al., 2004). Furthermore, there is a correlation between NMO-IgG titers and disease activity (Takahashi et al., 2007). It has been identified that NMO-IgG binds to the extracellular domains of AQP4 although it is not clear if the binding of NMOIgG to AQP4 alters the water permeability of AQP4 (Takahashi et al., 2006; Hinson et al., 2007). A marked loss of perivascular AQP4 immunoreactivity within NMO lesions is accompanied by intense vasculocentric immune complex deposition (Jarius et al., 2008). These findings suggest that a complement-activating, AQP4-specific NMO-IgG might initiate the NMO lesion rather than be a result of the disease. However, many aspects of the pathogenesis of NMO remain unclear. For example, it has to be evaluated how loss of AQP4 expression is related to the initiation and/or the progression of the disease.

7

Conclusion

Water transport is coupled to a number of brain functions as well as to pathological conditions such as brain edema, tumors, and seizures. AQP4 is a predominant mammalian aquaporin in the central nervous system. Interestingly, AQP4 is not expressed in any neurons but is exclusively expressed in astrocytes, where it is polarized in their processes in contact with vessels, ependymal cells and pia, suggesting a role in water movement into and out of brain parenchyma. The concept has emerged that astrocytes not only contribute to the clearance of transmitters and K+ from synaptic regions but that they also control the local extracellular space volume, where AQP4 plays a central role. AQP4 is also important in the pathophysiology of cytotoxic and vasogenic brain edema, although it has opposing roles. The seizure phenotype in AQP4-null mice suggests the possibility of AQP4 modulation in epilepsy therapy. The clinical relevance of AQP4 is largely unknown. However, the finding that NMO-IgG specifically binds to AQP4 in NMO patients gives us a clue regarding the role of AQP4 in human pathophysiology. Thus there are exciting possibilities for AQP4-based therapy in a variety of common disorders of the central nervous system.

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Aquaporins in the brain Nase G, Helm PJ, Enger R, Ottersen OP. 2008. Water entry into astrocytes during brain edema formation. Glia 56: 895-902. Neely JD, Christensen BM, Nielsen S, Agre P. 1999. Heterotetrameric composition of aquaporin-4 water channels. Biochemistry 38: 11156-11163. Neely JD, Amiry-Moghaddam M, Ottersen OP, Froehner SC, Agre P, et al. 2001. Syntrophin-dependent expression and localization of Aquaporin-4 water channel protein. Proc Natl Acad Sci USA 98: 14108-14113. Nesic O, Lee J, Ye Z, Unabia GC, Rafati D, et al. 2006. Acute and chronic changes in aquaporin 4 expression after spinal cord injury. Neuroscience 143: 779-792. Nicchia GP, Nico B, Camassa LM, Mola MG, Loh N, et al. 2004. The role of aquaporin-4 in the blood-brain barrier development and integrity: Studies in animal and cell culture models. Neuroscience 129: 935-945. Nielsen S, Nagelhus EA, Amiry-Moghaddam M, Bourque C, Agre P, et al. 1997. Specialized membrane domains for water transport in glial cells: High-resolution immunogold cytochemistry of aquaporin-4 in rat brain. J Neurosci 17: 171-180. Oshio K, Watanabe H, Song Y, Verkman AS, Manley GT. 2005. Reduced cerebrospinal fluid production and intracranial pressure in mice lacking choroid plexus water channel Aquaporin-1. Faseb J 19: 76-78. Padmawar P, Yao X, Bloch O, Manley GT, Verkman AS. 2005. K+ waves in brain cortex visualized using a long-wavelength K+-sensing fluorescent indicator. Nat Methods 2: 825-827. Papadopoulos MC, Verkman AS. 2005. Aquaporin-4 gene disruption in mice reduces brain swelling and mortality in pneumococcal meningitis. J Biol Chem 280: 13906-13912. Papadopoulos MC, Verkman AS. 2007. Aquaporin-4 and brain edema. Pediatric Nephrology (Berlin, Germany) 22: 778-784. Papadopoulos MC, Verkman AS. 2008. Potential utility of aquaporin modulators for therapy of brain disorders. Prog Brain Res 170: 589-601. Papadopoulos MC, Manley GT, Krishna S, Verkman AS. 2004. Aquaporin-4 facilitates reabsorption of excess fluid in vasogenic brain edema. Faseb J 18: 1291-1293. Preston GM, Agre P. 1991. Isolation of the cDNA for erythrocyte integral membrane protein of 28 kilodaltons: member of an ancient channel family. Proc Natl Acad Sci USA 88: 11110-11114. Preston GM, Carroll TP, Guggino WB, Agre P. 1992. Appearance of water channels in Xenopus oocytes expressing red cell CHIP28 protein. Science (New York, N.Y.) 256: 385-387. Preston GM, Jung JS, Guggino WB, Agre P. 1993. The mercury-sensitive residue at cysteine 189 in the CHIP28 water channel. J Biol Chem 268: 17-20.

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Rash JE, Yasumura T, Hudson CS, Agre P, Nielsen S. 1998. Direct immunogold labeling of aquaporin-4 in square arrays of astrocyte and ependymocyte plasma membranes in rat brain and spinal cord. Proc Natl Acad Sci USA 95: 11981-11986. Rite I, Machado A, Cano J, Venero JL. 2008. Intracerebral VEGF injection highly upregulates AQP4 mRNA and protein in the perivascular space and glia limitans externa. Neurochem Intl 52: 897-903. Rojek AM, Skowronski MT, Fuchtbauer EM, Fuchtbauer AC, Fenton RA, et al. 2007. Defective glycerol metabolism in aquaporin 9 (AQP9) knockout mice. Proc Natl Acad Sci USA 104: 3609-3614. Saadoun S, Papadopoulos MC, Davies DC, Krishna S, Bell BA. 2002. Aquaporin-4 expression is increased in oedematous human brain tumours. J Neurol Neurosurg Psychiatry 72: 262-265. Sui H, Han BG, Lee JK, Walian P, Jap BK. 2001. Structural basis of water-specific transport through the AQP1 water channel. Nature 414: 872-878. Suzuki H, Nishikawa K, Hiroaki Y, Fujiyoshi Y. 2008. Formation of aquaporin-4 arrays is inhibited by palmitoylation of N-terminal cysteine residues. Biochimica et Biophysica Acta 1778: 1181-1189. Tait MJ, Saadoun S, Bell BA, Papadopoulos MC. 2008. Water movements in the brain: Role of aquaporins. Trends Neurosci 31: 37-43. Takahashi T, Fujihara K, Nakashima I, Misu T, Miyazawa I, et al. 2006. Establishment of a new sensitive assay for antihuman aquaporin-4 antibody in neuromyelitis optica. Tohoku J Exp Med 210: 307-313. Takahashi T, Fujihara K, Nakashima I, Misu T, Miyazawa I, et al. 2007. Anti-aquaporin-4 antibody is involved in the pathogenesis of NMO: A study on antibody titre. Brain 130: 1235-1243. Takumi Y, Nagelhus EA, Eidet J, Matsubara A, Usami S, et al. 1998. Select types of supporting cell in the inner ear express aquaporin-4 water channel protein. Euro J Neurosci 10: 3584-3595. Vajda Z, Pedersen M, Fuchtbauer EM, Wertz K, StodkildeJorgensen H, et al. 2002. Delayed onset of brain edema and mislocalization of aquaporin-4 in dystrophinnull transgenic mice. Proc Natl Acad Sci USA 99: 13131-13136. Verbavatz JM, Ma T, Gobin R, Verkman AS. 1997. Absence of orthogonal arrays in kidney, brain and muscle from transgenic knockout mice lacking water channel aquaporin-4. J Cell Sci 110 (Pt 22): 2855-2860. Verkman AS, Binder DK, Bloch O, Auguste K, Papadopoulos MC. 2006. Three distinct roles of aquaporin-4 in brain function revealed by knockout mice. Biochimica et Biophysica Acta 1758: 1085-1093.

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Wakayama Y, Jimi T, Inoue M, Kojima H, Murahashi M, et al. 2002. Reduced aquaporin 4 expression in the muscle plasma membrane of patients with Duchenne muscular dystrophy. Arch Neurol 59: 431-437. Warth A, Simon P, Capper D, Goeppert B, Tabatabai G, et al. 2007. Expression pattern of the water channel aquaporin-4 in human gliomas is associated with blood-brain barrier disturbance but not with patient survival. J Neurosci Res 85: 1336-1346. Wen H, Nagelhus EA, Amiry-Moghaddam M, Agre P, Ottersen OP, et al. 1999. Ontogeny of water transport in rat brain: Postnatal expression of the aquaporin-4 water channel. Euro J Neurosci 11: 935-945. Wingerchuk DM, Lennon VA, Lucchinetti CF, Pittock SJ, Weinshenker BG. 2007. The spectrum of neuromyelitis optica. Lancet Neurol 6: 805-815.

Yang B, Brown D, Verkman AS. 1996. The mercurial insensitive water channel (AQP-4) forms orthogonal arrays in stably transfected Chinese hamster ovary cells. J Biol Chem 271: 4577-4580. Yukutake Y, Tsuji S, Hirano Y, Adachi T, Takahashi T, et al. 2008. Mercury chloride decreases the water permeability of aquaporin-4-reconstituted proteoliposomes. Biol Cell/ Under the Auspices of the Euro Cell Biol Organiz 100: 355-363. Zador Z, Magzoub M, Jin S, Manley GT, Papadopoulos MC, et al. 2008. Microfiberoptic fluorescence photobleaching reveals size-dependent macromolecule diffusion in extracellular space deep in brain. Faseb J 22: 870-879. Zelenina M, Zelenin S, Bondar AA, Brismar H, Aperia A. 2002. Water permeability of aquaporin-4 is decreased by protein kinase C and dopamine. Am J Physiol 283: F309-318.

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Voltage-Gated Calcium Ion Channels and Novel Voltage Sensing Proteins

Y. Okamura

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 406

2 2.1 2.1.1 2.1.2 2.1.3 2.2 2.2.1 2.2.2 2.2.3 2.2.4 2.3 2.4

Classifications of Voltage-Gated Ion Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 407 Neural Voltage-Gated Sodium Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 407 Nav1.1–1.3, Nav1.6, Nav1.7 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 407 Nav1.8, Nav1.9, Nav1.5 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 408 Nax . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 408 Neural Voltage-Gated Potassium Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 408 Kv1–Kv4 Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 408 KCNQ2/3 Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 409 HERG Potassium Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 409 BK Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 409 HCN Channel . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 409 Cyclic Nuleotide-Gated (CNG) Channels and Transient Receptor Potential (TRP) Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 410

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Human Diseases Related to Voltage-Gated Ion Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 410

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Regulation of Voltage-Gated Ion Channels by Auxiliary Subunits . . . . . . . . . . . . . . . . . . . . . . . . . . . 410

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Operation Mechanisms and Atomic Structure of the Voltage Sensor . . . . . . . . . . . . . . . . . . . . . . . . . 410

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Voltage-Sensing Phosphatase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 411

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Voltage-Gated Proton Channel: Ion Channel Without Pore . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 412

8

Future Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 413

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2009 Springer Science+Business Media, LLC.

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Voltage-gated calcium ion channels and novel voltage sensing proteins

Abstract: Voltage-gated ion channels are the key membrane proteins in the neural cells that underlie electrical signals, such as generation and propagation of action potentials, transmitter release, and synaptic integration. Voltage-gated ion channels consist of depolarization-activated groups with rigid ion permeabilities to sodium, calcium, and potassium, and hyperpolarization-activated group (HCN channel) that is permeable both to sodium and potassium. Each family of voltage-gated ion channel includes several genes that show distinct patterns of cell-specific expression and distinct subcellular localizations. Voltage-gated ion channels share common structure with six transmembrane segments. The first four transmembrane segments constitute the voltage-sensor domain (VSD) and the C-terminal two transmembrane segments provide pore domain. Recently, two novel voltage-sensor containing proteins were identified. VSP, voltage-sensing phosphatase, has the VSD and the phosphatase domain and shows voltage-dependent phosphoinositide phosphatase activity. VSOP/Hv1, which only consists of the VSD, exhibits activities of the voltage-gated proton channel. These suggest that voltage-evoked signals in the nervous system depend not only on voltage-gated ion channels but also on novel mechanisms that were previously unappreciated. List of Abbreviations: Cav channel, voltage-gated calcium channel; EAG channel, Ether-a´-go-go channel; HCN channel, hyperpolarization-activated, cyclic nucleotide-modified channel; Kv channel, voltage-gated potassium channel; Nav channel, voltage-gated sodium channel; S4, segment 4; VSD, voltage-sensor domain

1

Introduction

Since the introduction of the concept of ‘‘ion channels’’ for propagation and generation of action potentials in the squid giant axon, voltage-gated ion channels have been intensively studied as basic elements of electrical signals in neurons and muscle cells. Intrinsic excitabilities of neurons depend on the density, distribution patterns, and biophysical properties of voltage-gated ion channels. Voltage-gated sodium (Nav) and potassium (Kv) channels play principal roles in generation and propagation of action potentials in axons. They also determine membrane properties for synaptic integration in cell soma and dendrites. Voltage-gated calcium channels (Cav channels) play critical roles in regulation of transmitter release at presynaptic terminals, dendritic calcium influx for synaptic plasticity, and calcium influx in cell soma for gene regulation and synaptic maturation. Hyperpolarization-activated and cyclic nucleotide-modified channels (HCN channels) are both permeable to sodium and potassium. Nonselective cation permeability and hyperpolarization-gated opening render the membrane potential slowly depolarized following action potentials, generating slow rhythmic patterns of firing. Nav, Cav, Kv, and HCN channels share basic homologous units: alpha subunits of Nav or Cav channel consist of four homologous repeats, whereas in voltage-gated potassium channel or HCN channel, individual repeat is encoded by single gene and four subunits must assemble to form channels. The pore domain provides a narrow ionic pathway called selectivity filter that determines ion selectivity. Structure of bacterial potassium channel, KcsA, was defined by crystallography at atomic level and mechanisms of potassium-selective permeation are deeply understood (Zhou and MacKinnon, 2003). The voltage-sensor domain (VSD) senses transmembrane potential of less than 10 mV and translates this voltage change into remarkable increase of open probability of channel pore. This step is called voltagedependent gating. Voltage-gated sodium channels have steep voltage dependency and high sodium selectivity with positive equilibrium potential determined by sodium. These properties enable regenerative change of membrane potential, called ‘‘action potential.’’ Recent crystallographic studies (Long et al., 2005) of voltage-gated potassium channel indicated that the VSD is localized at the periphery of the channel, and parts of the voltage sensor are facing the lipid membranes. The operation of voltage sensing depends on signature pattern of amino acid alignments in the fourth segment (S4). Four to seven positively charged residues periodically align with two intervening hydrophobic residues. In response to membrane potential change, displacement of these charges occurs in the cell membrane that can be readout as the ‘‘gating currents.’’ In HCN channel, the coupling between the voltage sensor and the pore domain is reversed: the

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depolarization-induced movement of the VSD closes, not opens, ion channel gate. In a recently identified voltage-sensing protein, voltage-sensing phosphatase (VSP), phosphatase, instead of ion channel pore, is regulated by the VSD. These indicate that VSD is a transferable functional module. Since voltage-gated ion channels are the critical determinants of electrical properties of neurons combined with geometry and size of neurons, they are the important targets for modulation of excitabilities and synaptic transmission. Most types of voltage-gated ion channels are modulated by second messengers such as calcium, cAMP, G-proteins, and protein kinase C. Modulation of their activities is also mediated by other molecular mechanisms, including modulation by auxiliary subunits, calmodulin–Ca complex, and binding to phosphoinositides. Some voltage-gated ion channels are associated with small subunits or cytoplasmic proteins that have enzyme activities. One such example is modulation of Kv channel activities by beta subunit: activities of Kv channel are modified by the enzyme activities of Kv-beta subunit upon substrate binding (Weng et al., 2006). This chapter excludes voltage-dependent Cl channel, since voltage sensitivity of this type of channel is mediated by a distinct mechanism than that for already-mentioned voltage-sensitive channels.

2

Classifications of Voltage-Gated Ion Channels

2.1 Neural Voltage-Gated Sodium Channels Nav channels show sodium-selective permeability and rapid gating of activation and inactivation, playing key roles in initiation and propagation of action potentials in axons. Nav channels in cell soma and dendrite contribute to integration of synaptic inputs and retrograde propagation of action potentials into postsynaptic regions (this is called ‘‘back-propagation’’). In addition to rapidly inactivating sodium current, neuronal Nav current has subthreshold inward currents that contribute to repetitive firing and pacemaking. Steady-state current, often called ‘‘persistent sodium current’’ (Ina-p), is activated at subthreshold level. Exact molecular basis for Ina-p remains completely unknown. It is likely that Ina-p is derived from a modified mode of opening of fast-inactivating sodium channel. Second, many types of neurons show sodium current that has escaped from inactivation and reactivated following action potentials (called ‘‘resurgent sodium current’’ or Ina-res). Both Ina-p and Ina-res currents are activated at subthreshold levels thereby having large impacts on firing behaviors of neurons. Voltage-dependent gating depends on a S4 segment with positive charges in each homologous repeat. The S4 segment of the first two domains sense rapid depolarization, and then the other two segments contribute for gating of the pore. Voltage-gated sodium channels rapidly inactivate in a scale of 1–5 ms by a mechanism similar to the N-type inactivation of voltage-gated potassium channels: blocking particle provided from the part of the liner region between the repeat III and IV occludes the pore in a voltagedependent manner.

2.1.1 Nav1.1–1.3, Nav1.6, Nav1.7 These isoforms are TTX-sensitive fast-activated and inactivating sodium channels that are specifically expressed in the nervous system. Nav1.1–1.3 are mainly expressed in CNS neurons, whereas Nav1.7 is only expressed in peripheral neurons. Nav1.6 is both expressed in CNS and peripheral neurons. Nav1.6 is the major population of axonal sodium channels localized at node of Ranvier and initial segments that play a role in generation and propagation of action potentials. Nav1.6 is also expressed in cell soma in many types of neurons, contributing to ‘‘persistent sodium current’’ and ‘‘resurgent current.’’ Turnover of Nav channel isoforms occurs during maturation of myelinated nerve fibers. Nav1.2 is expressed at the node of Ranvier and replaced by Nav1.6 during maturation. Nav1.7 conducts many modalities, including pain, of sensory signals in peripheral nerves.

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2.1.2 Nav1.8, Nav1.9, Nav1.5 Nav1.8 and Nav1.9 are TTX resistant, slowly activating sodium channels that are exclusively expressed in DRG neurons. They play role in conduction of pain sensation. Nav1.8 and Nav1.9 have distinct voltage threshold of activation, kinetics, voltage sensitivity of inactivation, and temperature dependence. Nav1.8 plays a more important role in pain sensation at lower temperature, since null mouse of Nav1.8 exhibits defect of pain sensation at low temperature (Zimmermann et al., 2007). Nav1.5 is the TTX-resistant sodium channel that is predominantly expressed in cardiac cells. Nav1.5 is also expressed in the neurons of the limbic system. Physiological significance of this class of sodium channel in the limbic system still remains unknown.

2.1.3 Nax Nax has a primary structure with overall homology to mammalian voltage-gated sodium channels. However, basic amino acids in the S4 segment critical for voltage-dependent activation are not well conserved, and the III–IV linker critical for fast inactivation is also diverged. Recently, Nax has been shown to be expressed in glial cells of subfornical organ to play an unique function as an extracellular sodium sensor (Hiyama et al., 2002; Shimizu et al., 2007), but does not exhibit voltage-gated properties.

2.2 Neural Voltage-Gated Potassium Channels Kv channels negatively regulate neuronal excitabilities by damping membrane potentials toward equilibrium potentials for potassium. Modulation of Kv channels by ion concentration change or by second messengers caused by transmitter or hormone underlies plastic change of neuronal excitabilities. Kv channels share basic molecular structure with voltage-gated sodium and calcium channels; four transmembrane segments form the voltage sensor and the C-terminal two transmembrane segments provide the pore. In contrast with Nav channels and Cav channels, Kv channels consist of homologous or heterologous tetramers. For this assembly, different types of Kv channels contain distinct motifs for protein interactions in cytoplasmic regions. In addition, heterologous subunit assembly in Kv channels and HCN channels makes their functional diversities larger than Nav channels and Cav channels. Among voltage-gated ion channels, Kv channels have been best studied in terms of structure–function relationship because of its earlier identification of molecular species in the history and the presence of prokaryotic molecular homologs more suitable for crystallization. X-ray crystal structures of Kv channels under open state have been resolved (Long et al., 2005, 2007). In all classes of Kv channels, S4-segment with positively charged residues mediate voltage-dependent gating with distinct ranges of threshold and activation speed. Voltage-dependent gating is modified by binding of calcium to an intracellular side in large-conductance voltage and calcium-activated potassium channel (BK channel). Diverse Kv channels show distinct temporal profiles of activities and contribute to distinct phases during action potentials and intervals. Such diversity of Kv channel activities depends on multiple mechanisms of self-inhibition, so-called ‘‘inactivation’’: cytoplasmic structure occludes the pore in a voltage-dependent manner like a ball-and-chain mechanism in the Kv1-subfamily of Kv channels, and in another case, pore structure is modified in a voltage-dependent manner to be nonconducting state.

2.2.1 Kv1–Kv4 Channels These belong to authentic Kv channels, which are closely related to Drosophila Shaker-type potassium channel, one of the best characterized ion channels. These four subfamilies show distinct kinetics of activation and inactivation and modify shapes of action potentials in distinct manner. Distinction of the

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four subfamilies is conserved from Drosophila to human, and they do not form heteromeric channels over members from a different subfamily. Kv1–Kv4 channels show distinct subcellular localizations and have distinct roles in transmitter release, spike initiation, synaptic integration at dendrites, and rhythmic firing patterns. Potassium currents mediated by Kv1–Kv4 channels are grouped into two types of currents with distinct biophysical properties: noninactivating or slowly inactivating K current and transient outward current (IA current). The former type is found in all neurons and present in cell soma and dendrite. The K current encoded by Kv3 shows extremely rapid activation and plays an important role in action potentials with high frequency. IA current showing rapid inactivation is present in dendrite, axon, and cell soma, playing an important role in determining threshold, latency, and frequency of neuronal firing.

2.2.2 KCNQ2/3 Channels M-current is the potassium conductance that was classically described in sympathetic neurons and hippocampal neurons as the current suppressed by muscarinic receptor stimulation (‘‘M’’ or M-current is in fact derived from the initial of muscarine). M-current has an important role in tuning neuronal excitabilities for its activities at resting membrane potentials and susceptibility to modulation through numerous receptors. The molecular correlate for the M-current is the heteromeric channels that consist of KCNQ2 and KCNQ3 (or KCNQ4 or KCNQ5). Heteromeric channels consisting of KCNQ2 and the other subunits are sensitive to PtdIns(4,5)P2. Hydrolysis of PtdIns(4,5)P2 upon activation of muscarnic acetylcholine receptor leads to reduction of M-current. KCNQ2 and KCNQ3 are also the dominant population of potassium channels in the axons: they are coclustered with Nav channels in the node of Ranvier and initial segments of myelinated nerves, regulating firing and propagation of action potentials.

2.2.3 HERG Potassium Channels HERG channels are rapidly activating potassium channels and are known to play critical roles in rapid repolarization following action potentials in cardiac cells. HERG channels are only expressed in glial cells but not in neurons, and potentially play roles in potassium homeostasis in the brain.

2.2.4 BK Channels BKCa channels are large-conductance voltage and calcium-activated potassium channel. BKCa channels are expressed in most types of neurons, and control cell excitability and neurotransmitter release, coupling the membrane potential with intracellular Ca2+ levels. BKCa channels have both calcium sensor and voltage sensor, and either stimulation of increase of intracellular calcium or depolarization increases the sensitivity to the other modality, thereby exhibiting synergism of activation. BK channels are often colocalized with Cav channels at presynaptic terminals. Ca influx through Cav channels quickly activates BK channels in the presynaptic terminal, sharpening the action potential and restricting time window of local Ca influx for vesicle release through Cav channels. Intrinsic Ca sensitivity of BK channel is based on the two sites of the cytoplasmic region that have distinct binding affinity for Ca ions. Interaction between the two Ca binding sites makes wide dynamic range of Ca sensitivity.

2.3 HCN Channel HCN channel is a cation channel permeable to both sodium and potassium. HCN channels are expressed in dendrites and cell soma. They play roles in forming pace-making activities through slowly depolarizing membrane potentials. HCN channels are also expressed in dendrites and important for synaptic integration.

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2.4 Cyclic Nuleotide-Gated (CNG) Channels and Transient Receptor Potential (TRP) Channels CNG channels and TRP channels have the transmembrane domain corresponding to the voltage sensor of authentic voltage-gated ion channels. However, signature alignment pattern of positive charges in S4 is not well conserved in these channels. Whether these channels have intrinsic voltage sensitivity is still controversial.

3

Human Diseases Related to Voltage-Gated Ion Channels

Genetic defects of neurally expressed voltage-gated ion channels lead to hereditary disorders with symptoms of abnormal neuronal excitabilities, including epilepsy, myoclony, and migraine. Mutations of alpha subunits (Nav1.1, Nav1.2) or beta subunit (Nav-beta1) of voltage-gated sodium channels and KCNQ2 or KCNQ3 gene cause epilepsy. Mutations in the gene encoding Kv1.1, a voltage-gated neuronal potassium channel, are associated with the disorder, called episodic ataxia type 1, which is a paroxysmal neurological disorder characterized by short-lived attacks of recurrent midline cerebellar dysfunction and continuous motor activity. Mutations of Nav1.7 gene that is exclusively expressed in the peripheral nervous system have recently been found in members that exhibit a congenital inability to experience pain (Cox et al., 2006). Human genetic disorders of voltage-gated calcium channels are described in Chapter xx.

4

Regulation of Voltage-Gated Ion Channels by Auxiliary Subunits

As in many other ion channel families, voltage-gated channels are regulated by auxiliary subunits. In voltage-gated sodium channels, localization and kinetics of alpha subunits are modulated by beta subunits. Among Nav-beta subunits, Beta4 subunit has a unique property of modulating channel functions: it plugs a pore of alpha subunits upon a short depolarization. During mild hyperpolarization following this depolarization, it is released from the pore, causing transient inward flow of sodium ions that underlie ‘‘resurgent sodium current.’’ Following this reopening of channel, sodium channel pore is plugged again by an intrinsic inactivation ball that corresponds to the III–IV linker region of the alpha subunit (Grieco et al., 2005). ‘‘Resurgent sodium current’’ contributes to repetitive firing of many types of CNS neurons including cerebellar Purkinje neurons and other neurons. One species of auxiliary subunit of Kv channel, Kv-beta, is another example that modifies channel kinetics by operating as the blocking particle. Some Kv-beta subunits include an amino-terminal region that allows them to transform non-inactivating Kv1 channels into rapidly inactivating channels. In addition, another Kv beta subunit, one for Shaker-or-Kv1 family potassium channel (Kv1), has another role: aldoketo-reductase activity that modulates the amplitude and kinetics of the alpha subunit of potassium channel (Kv1) in response to redox state in the cell (Weng et al., 2006). The beta subunit of the large conductance Ca(2+) and voltage-activated K(+) channel (BK(Ca)) modulates the apparent Ca(2+)/ voltage sensitivity, pharmacological and kinetic properties of the alpha subunit. Auxiliary subunits do not only regulate kinetics of the main subunits, but also regulate expression of channels to the cell surface. A cytoplasmic stretch of alpha subunit of Cav channel includes ER-retention signal, thereby making the whole protein pooled in the ER. Cav-beta subunit binds to this stretch to mask ER-retention signal, facilitating expression of the whole Cav channel complexes to the cell membrane.

5

Operation Mechanisms and Atomic Structure of the Voltage Sensor

The VSD of 6TM-type voltage-gated channels consists of four transmembrane helices as the VSD and the pore domain. The pore domain consists of two transmembrane segments with a short middle loop that forms a narrow ion pathway, called a ‘‘selectivity filter.’’ Selectivity filter determines ion selectivity and

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conductance. In the VSD, the fourth transmembrane segment, called S4, has signature sequence in which positively charged residues are periodically aligned with intervening hydrophobic residues. Mutagenesis studies suggested that these positive charges play a critical role in voltage sensing (Aggarwal and MacKinnon, 1996). Direct proof that S4 moves in response to change of membrane potential, and that positive charges in S4 indeed carry charges of gating currents, was obtained by extensive studies that detected changes in the accessibilities to methanethiosulfonate (MTS) reagents (Yang and Horn, 1995) and protons (Starace and Bezanilla, 2001), movement of fluorescent label of the substituted cysteine residues in S4 (Mannuzzu and Isacoff, 2000), or changes in current kinetics upon site-specific photo-cross-linking (Bezanilla, 2000; Horn et al., 2000). This structure of S4 is unusual as transmembrane segment, since it has hydrophilic residues. In these days, much attention by biophysicists has been made to the question how it is located in the cell membrane and how S4 orients to sense change of transmembrane voltage. Recent findings strongly support the idea that S1–S4 is a modular unit for voltage sensing and relatively free-moving unit slightly isolated from the pore domain. First, crystallization of mammalian Kv1.2 channel indicated that VSD, consisting of S1–S4, exists in the external core of the channel, and positively charged residues of S4 face the lipid (Long et al., 2005). Second, a recently discovered protein, VSP, contains VSD and phosphatase domain, in which the VSD regulates activity of enzyme (Murata et al., 2005). Three main models have been proposed for conformational change of the voltage sensor. In the canonical model, S4 is located in a hole, surrounded by other transmembrane helices and slide perpendicularly through the hole in response to membrane potential change. In the paddle model, S4 is facing the lipid and transverses the membrane with a large motion like a paddle. In the transporter model, motion of S4 is not large, and focused electric field changes by tilt of transmembrane helices either with motion of S4 or coordinated motion of other transmembrane helices. Crystal structures of Kv channel at open state were obtained from prokaryotic potassium channel, KvAP as the isolated VSD or protein complex with antibody (Jiang et al., 2003), and later, mammalian Kv1.2 was crystallized and resolved as a complex with Kv-beta subunit (Long et al., 2005). These results pictured that the VSD is located outside of the pore domain, facing the lipid. Recently, an open state structure of the voltage sensor of Kv channels was obtained with better resolution (2.4A˚) from a chimeric protein between Kv1.2 and Kv2.1 (Long et al., 2007), revealing that negative charges in S1–S3 form salt bridges, with positive charges of S4 playing an important role in the stabilization of voltage sensor. Mechanisms of coupling between VSD and pore domain remain unclear. Two mechanisms have been proposed based on the structure of Kv channels and biophysical studies. A linker region between S4 and pore domain has been proposed to pull the pore domain in response to voltage sensor movement and induces the conformational change of pore structure. The crystal structure of Kv1.2 channel (Long et al., 2007) is consistent with this model. On the other hand, VSD is apposed to the pore domain with a part of S4 facing helices of the pore domain. This structure suggests that movement is transmitted from VSD to pore domain through interactions among transmembrane helices, like a wheel.

6

Voltage-Sensing Phosphatase

Bioinformatic approach led to identification of Ci-VSP, ‘‘Ciona intestinalis voltage-sensor containing phosphatase,’’ which contains four transmembrane segments with significant homology to VSD, but lacked a pore domain (Murata et al., 2005) (> Figure 22-1). The C terminus shows homology to the wellcharacterized phosphatase PTEN (phosphatase and tensin homologue deleted on chromosome 10) (Maehama et al., 2001). The VSD of Ci-VSP, consisting of four transmembrane segments, shows displacement currents in response to membrane potential change when expressed in Xenopus oocytes, indicating that VSD operates as the voltage sensor. The cytoplasmic region of Ci-VSP shows significant homology to PTEN and has a CX5R motif that is conserved among PTEN, other phosphoinositide phosphatases (Maehama et al., 2001), protein tyrosine phosphatase (PTP), and other related phosphatases. The purified recombinant cytoplasmic region of Ci-VSP has the activity of dephosphorylation with phosphatidylinositol-3,4,5-trisphosphate (PtdIns(3,4,5)P3) as substrate (Murata et al., 2005), and this activity depends on cysteine residue in the CX5R consensus motif of the active center, as in many PTPs and PTEN.

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. Figure 22-1 Membrane topology of voltage-gated ion channels and voltage-sensor domain proteins. Voltage-gated ion channels consist of six transmembrane segments. The first four transmembrane segments constitute the voltage-sensor domain. The C-terminal two transmembrane segments constitute the pore domain. In voltage-sensing phosphatase (VSP), pore domain is missing and instead, phosphatase domain is attached downstream of the VSD. VSOP (voltage-sensor only protein) exhibits activities of voltage-gated proton channel

Ci-VSP is depolarization-activated phosphatidylinositol-4,5-bis-phosphate (PtdIns(4,5)P2) phosphatase. Its voltage-dependent enzyme activity was shown by experiments using Kir channels (Huang et al., 1998) and KCNQ potassium channels (Zhang et al., 2003) that are sensitive to phosphatidylinositol-3,4-bisphosphate (PtdIns(4,5)P2). Confocal imaging experiments using GFP-conjugated PH-domain derived from PLC delta subunit as the sensor for PtdIns(4,5)P2 indicated that PtdIns(4,5)P2 concentration is increased during hyperpolarization and decreased during depolarization(Murata and Okamura, 2007). Ci-VSP-like gene is conserved from sea urchin to humans, but is not found in the genomes of Drosophila or C. elegans. Ci-VSP is weakly expressed in the nervous system (Murata et al., 2005). The discovery of Ci-VSP suggests that voltage sensing is not restricted to functions of ion channels but more widely used than previously thought.

7

Voltage-Gated Proton Channel: Ion Channel Without Pore

Voltage-gated proton channels (Hv channels) were first described as the outward currents that induce intracellular alkalization in snail neurons. Later, similar currents were identified from amphibian oocytes, mammalian blood cells using whole cell patch recording. The Hv channel has been best characterized in the plasma membranes of blood cells including macrophage, neutrophils, and eosinophils that undergo phagocytosis. In the brain, Hv channels are expressed in microglia and potentially play roles in regulating extracellular pH at extracellular environments and/or activities of phagocytosis of microglia. Hv channels contain several unique biophysical properties. First, Hv channels are perfectly selective to protons over other cations. Hv channels show extremely slow activation kinetics that becomes faster with depolarization. This voltage dependency is pH dependent. The activation threshold is more positive when

Voltage-gated calcium ion channels and novel voltage sensing proteins

22

delta pH (pHi pHo) is larger. With this property, proton current is always activated at the voltage range where outward proton current flows. Hv channels show sensitivity to micromolar zinc and to higher doses of cadmium (Decoursey, 2003). The Hv channel in mammalian phagocytes was originally proposed as the proton-transporting pathway that regulates intracellular pH during oxygen consumption associated with phagocytosis, which is called ‘‘respiratory burst’’ (Henderson et al., 1987). Each cycle of NADPH oxidase enzyme activity leaves two protons in the intracellular side (Murphy and DeCoursey, 2006). Accumulation of protons by NADPH oxidase activity causes large charge imbalance, leading to rapid depolarization and acidification in the cytoplasm. Since the Hv channel has the nature of activation both by depolarization and intracellular acidification, NADPH oxidase activity efficiency activates Hv channel. Physiological roles of Hv channel in phagocyte are thought to be several-fold. First, Hv channel is the major pathway that compensates for charge imbalance that results from the efflux of electron due to NADPH oxidase activities. NADPH oxidase activity is known to be attenuated at potentials more than 50 mV. Proton flux through Hv channel prevents the membrane potential from becoming too positive so that NADPH oxidase activities are kept active (Decoursey, 2003). In fact, submillimolar zinc that is sufficient to silence the Hv current significantly reduces production rate of superoxide anion in human eosinophils, probably through depolarizationinduced inhibition of NADPH oxidase activity (DeCoursey et al., 2003). Second, activities of the Hv channel also maintain intracellular pH neutral to keep ROS generation, since efficiency of NADPH oxidase activities decrease by low intracellular pH (Clark et al., 1987; Morgan et al., 2005). Third, Hv channels not only regulate pH and electro-neutrality in cytoplasm, but could also provide protons in phagosome, which is a closed membrane compartment. A gene that encodes voltage-gated proton current has recently been identified as a protein (VSOP or Hv1) that contains four transmembrane segments with significant homology to VSD of voltage-gated ion channels. Surprisingly, this protein lacks pore domain (Sasaki et al., 2006). Its S4-like segment has three arginines separated by two hydrophobic residues as in 6-TM voltage-gated channels, although the number of positively charged residues is few in this protein. Despite the lack of pore domain, heterologously expressed VSOP/Hv1 proteins in mammalian cells and Xenopus oocyte recapitulated all behaviors of native Hv currents. How could protons permeate despite absence of apparent pore? In several examples of proton-conducting proteins, protons are transferred by hopping through hydrogen-bonded chains rather than by ion diffusion through an aqueous pore. In that case, individual protons do not need to physically pass all the way through the channel pathway. On the other hand, the proton-permeating activities of this protein are reminiscent of phenotypes of histidine-replacing mutants in S4 of Shaker potassium channel (Starace and Bezanilla, 2001, 2004). Since histidine has a pKa near pH 7 and the charge on the residue could be regulated simply by pH, accessibility of proton to histidine-replaced residues could report the conformation of S4 and its environmental electric field (Starace and Bezanilla, 2001, 2004). Some mutants show pH-dependent change of gating current, whereas other mutants, including R371H, show outward or inward voltagedependent proton conductances (Starace and Bezanilla, 2001). A finding that VSD acquires ion permeation properties by mutation was also reported in other mutants of Kv channel (Sokolov et al., 2005) and Nav channels (Sokolov et al., 2005). Mutants replacing arginines by smaller, uncharged residues (Ala, Ser, Cys, Val) show permeability to potassium, sodium, and cesium during hyperpolarization (Tombola et al., 2005), being called ‘‘omega current.’’ In addition, human Nav1.4 sodium channel from patients of hypokalemic periodic paralysis, which has mutation in the VSD, shows nonselective ion permeation (Sokolov et al., 2007) or proton permeation (Struyk and Cannon, 2007). All these findings suggest that VSD itself has a potentiality of ion permeation. At present, it is unclear whether voltage-gated proton channels share mechanisms of proton conduction with mutated VSDs of conventional voltage-gated ion channels.

8

Future Perspectives

From 1980 to 2000, the majority of molecular species of ion channels that underlie neuronal excitabilities have been identified, and robust studies have been done for their structure–function relationships. In

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contrast, correlations between molecules and native currents have not been completely established. For example, KCNQ2/3 originally identified as the molecular correlate for M-current has been rediscovered as the important potassium channels that are clustered at nodes of myelinated nerve fibers and play critical roles in action potential generation and propagation (Pan et al., 2006). Moreover, unexpected roles of voltage-gated ion channels have been emerging. Nax is a homolog of voltage-gated sodium channel, which is mainly expressed in glial cells. Nax does not operate as the Nav channel but functions as a sensor for extracellular sodium concentration (Hiyama et al., 2002) for sodium intake behavior (Shimizu et al., 2007). Some voltage-gated ion channels have non-ion-conducting functions. Drosophila ether-a´-go-go channel (EAG) channels operate as action potential dumper by repolarizing membrane potentials, but the nonconductive mutant channels can still trigger intracellular chemical signal for cell proliferation without being accompanied by potassium flow (Hegle et al., 2006). Although these mechanisms still remain unknown, such recent findings of ion channels may be relevant to a well-accepted idea of skeletal muscle L-type Cav channel that transduces electrical signal into contraction through directly coupling to ryanodine receptors, intracellular Ca channels, without requiring Ca permeability. Some auxiliary subunits for voltage-gated ion channels operate as enzymes as shown in the aldo-keto-reductase activity of Kv-beta subunit (Weng et al., 2006). These raise a possibility that the conformational change of voltage sensor directly regulates enzyme activity of proteins that are associated with the main channel subunit. In fact, the discovery of Ci-VSP has brought the idea that voltage sensor operates not only for ion pore but also for other cellular functions. Therefore, electro-chemical signaling in cell membranes could be based on more diverse molecular mechanisms than that previously thought.

References Aggarwal SK, MacKinnon R. 1996. Contribution of the S4 segment to gating charge in the Shaker K+ channel. Neuron 16: 1169-1177. Bezanilla F. 2000. The voltage sensor in voltage-dependent ion channels. Physiol Rev 80: 555-592. Clark RA, Leidal KG, Pearson DW, Nauseef WM. 1987. NADPH oxidase of human neutrophils. Subcellular localization and characterization of an arachidonateactivatable superoxide-generating system. J Biol Chem 262: 4065-4074. Cox JJ, Reimann F, Nicholas AK, Thornton G, Roberts E, et al. 2006. An SCN9A channelopathy causes congenital inability to experience pain. Nature 444: 894-898. Decoursey TE. 2003. Voltage-gated proton channels and other proton transfer pathways. Physiol Rev 83: 475-579. DeCoursey TE, Morgan D, Cherny VV. 2003. The voltage dependence of NADPH oxidase reveals why phagocytes need proton channels. Nature 422: 531-534. Grieco TM, Malhotra JD, Chen C, Isom LL, Raman IM. 2005. Open-channel block by the cytoplasmic tail of sodium channel beta4 as a mechanism for resurgent sodium current. Neuron 45: 233-244. Hegle AP, Marble DD, Wilson GF. 2006. A voltage-driven switch for ion-independent signaling by ether-a-go-go K+ channels. Proc Natl Acad Sci USA 103: 2886-2891. Henderson LM, Chappell JB, Jones OT. 1987. The superoxidegenerating NADPH oxidase of human neutrophils is

electrogenic and associated with an H+ channel. Biochem J 246: 325-329. Hiyama TY, Watanabe E, Ono K, Inenaga K, Tamkun MM, et al. 2002. Na(x) channel involved in CNS sodium-level sensing. Nat Neurosci 5: 511-512. Horn R, Ding S, Gruber HJ. 2000. Immobilizing the moving parts of voltage-gated ion channels. J Gen Physiol 116: 461-476. Huang CL, Feng S, Hilgemann DW. 1998. Direct activation of inward rectifier potassium channels by PIP2 and its stabilization by Gbg. Nature 391: 803-806. Jiang Y, Lee A, Chen J, Ruta V, Cadene M, et al. 2003. X-ray structure of a voltage-dependent K+ channel. Nature 423: 33-41. Long SB, Campbell EB, Mackinnon R. 2005. Crystal structure of a mammalian voltage-dependent Shaker family K+ channel. Science 309: 897-903. Long SB, Tao X, Campbell EB, MacKinnon R. 2007. Atomic structure of a voltage-dependent K+ channel in a lipid membrane-like environment. Nature 450: 376-382. Maehama T, Taylor GS, Dixon JE. 2001. PTEN and myotubularin: Novel phosphoinositide phosphatases. Annu Rev Biochem 70: 247-279. Mannuzzu LM, Isacoff EY. 2000. Independence and cooperativity in rearrangements of a potassium channel voltage sensor revealed by single subunit fluorescence. J Gen Physiol 115: 257-268.

Voltage-gated calcium ion channels and novel voltage sensing proteins Morgan D, Cherny VV, Murphy R, Katz BZ, DeCoursey TE. 2005. The pH dependence of NADPH oxidase in human eosinophils. J Physiol 569: 419-431. Murata Y, Iwasaki H, Sasaki M, Inaba K, Okamura Y. 2005. Phosphoinositide phosphatase activity coupled to an intrinsic voltage sensor. Nature 435: 1239-1243. Murata Y, Okamura Y. 2007. Depolarization activates the phosphoinositide phosphatase Ci-VSP, as detected in Xenopus oocytes coexpressing sensors of PIP2. J Physiol 583: 875–889. Murphy R, DeCoursey TE. 2006. Charge compensation during the phagocyte respiratory burst. Biochim Biophys Acta 1757: 996-1011. Pan Z, Kao T, Horvath Z, Lemos J, Sul JY, et al. 2006. A common ankyrin-G-based mechanism retains KCNQ and NaV channels at electrically active domains of the axon. J Neurosci 26: 2599-2613. Sasaki M, Takagi M, Okamura Y. 2006. A voltage sensordomain protein is a voltage-gated proton channel. Science 312: 589-592. Shimizu H, Watanabe E, Hiyama TY, Nagakura A, Fujikawa A, et al. 2007. Glial Nax channels control lactate signaling to neurons for brain [Na+] sensing. Neuron 54: 59-72. Sokolov S, Scheuer T, Catterall WA. 2005. Ion permeation through a voltage-sensitive gating pore in brain sodium channels having voltage sensor mutations. Neuron 47: 183-189. Sokolov S, Scheuer T, Catterall WA. 2007. Gating pore current in an inherited ion channelopathy. Nature 446: 76-78.

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Starace DM, Bezanilla F. 2001. Histidine scanning mutagenesis of basic residues of the S4 segment of the shaker K+ channel. J Gen Physiol 117: 469-490. Starace DM, Bezanilla F. 2004. A proton pore in a potassium channel voltage sensor reveals a focused electric field. Nature 427: 548-553. Struyk AF, Cannon SC. 2007. A Na+ channel mutation linked to hypokalemic periodic paralysis exposes a proton-selective gating pore. J Gen Physiol 130: 11-20. Tombola F, Pathak MM, Isacoff EY. 2005. Voltage-sensing arginines in a potassium channel permeate and occlude cation-selective pores. Neuron 45: 379-388. Weng J, Cao Y, Moss N, Zhou M. 2006. Modulation of voltage-dependent Shaker family potassium channels by an aldo-keto reductase. J Biol Chem 281: 15194-15200. Yang N, Horn R. 1995. Evidence for voltage-dependent S4 movement in sodium channels. Neuron 15: 213-218. Zhang H, Craciun LC, Mirshahi T, Rohacs T, Lopes CM, et al. 2003. PIP2 activates KCNQ channels, and its hydrolysis underlies receptor-mediated inhibition of M currents. Neuron 37: 963-975. Zhou Y, MacKinnon R. 2003. The occupancy of ions in the K+ selectivity filter: Charge balance and coupling of ion binding to a protein conformational change underlie high conduction rates. J Mol Biol 333: 965-975. Zimmermann K, Leffler A, Babes A, Cendan CM, Carr RW, et al. 2007. Sensory neuron sodium channel Nav1.8 is essential for pain at low temperatures. Nature 447: 855-858.

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Muscarinic Acetylcholine Receptor

S. Ichiyama . T. Haga

1 1.1 1.2 1.3

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 418 History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 418 Distribution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 419 Diseases and Therapeutic Agents Related to Muscarinic Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 420

2 2.1 2.2 2.3 2.4 2.5 2.5.1 2.5.2 2.5.3

Structure and Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 420 Structural Features . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 420 Agonist Binding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 420 Coupling to G Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 422 Conformational Change upon Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 424 Regulation of Effector Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 426 Activation of Phospholipase C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 426 Regulation of Adenylyl Cyclase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 426 Regulation of Ion Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427

3 3.1 3.2 3.3

Regulation of Receptor Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427 Desensitization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427 Phosphorylation by GRK and Receptor Internalization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 428 Regulation by RGS Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 428

4

Dimerization/Oligomerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 429

5

Complex Formation with Other Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 429

6 6.1 6.2 6.3 6.4 6.5

Knockout Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 430 M1 KO Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 430 M2 KO Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 430 M3 KO Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 430 M4 KO Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 431 M5 KO Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 431

7

Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 431

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2009 Springer Science+Business Media, LLC.

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Muscarinic acetylcholine receptor

Abstract: Muscarinic acetylcholine receptors (muscarinic receptors) are prototypical G protein-coupled receptors (GPCRs) activated by the endogenous neurotransmitter acetylcholine and play key roles in regulating the activity of many vital functions of the central and peripheral nervous systems. Studies of muscarinic receptors as well as rhodopsin and b adrenergic receptors have revealed several characteristics of GPCRs, which are also true for other GPCRs. Here, we present an overview of the historical background and current studies of muscarinic receptors focusing on studies carried out in our laboratory. List of Abbreviations: AC, adenylyl cyclase; AKAP, cAMP-dependent protein kinase anchoring protein; GAP, GTPase-activating protein; GPCR, G protein-coupled receptor; GRK, G protein-coupled receptor kinase; i2, second intracellular loop; i3, third intracellular loop; IP3, inositol-1,4,5-trisphosphate; KO, knockout; NMR, nuclear magnetic resonance; PKA, cAMP-dependent protein kinase; PKC, protein kinase C; PLC, phospholipase C; RGS, regulators of G protein signaling; RH, regulator of G protein signaling homology; TM, transmembrane; TRPC6, transient receptor potential-canonical subtype; WT, wild-type

1

Introduction

1.1 History The effects of acetylcholine can be divided into nicotinic and muscarinic based on differences in the effects of acetylcholine mimetics (agonists) and competitive inhibitors (antagonists) on diverse tissues. The term ‘‘muscarinic’’ comes from the fact that the effects of muscarine on the cells innervated by postganglionic parasympathetic neurons are similar to those caused by stimulation of the neurons. Muscarine is a natural alkaloid derived from the poisonous mushroom, Amanita muscaria, whose adverse effects include salivation, lacrimation, nausea, headache, visual disturbances, bradycardia, and hypotension (Brown and Taylor, 2001). The chemical structures of acetylcholine, other agonists, and antagonists are shown in > Figure 23-1. The concept of a ‘‘receptor’’ that exists on the cell surface, accepts acetylcholine, and triggers cell responses, had long been established before its molecular details were elucidated. The development of radiolabeled ligands made it possible to characterize the ligand-binding activities of receptors in intact tissues, intact cells, or membranes prepared from cells. The presence of several subtypes was suggested based on the heterogeneous binding properties of muscarinic antagonists, such as pirenzepine (Hammer et al., 1980). The specific activity of ligand binding to receptors in membrane preparations is rarely in excess of 1 pmol/ mg protein, indicating that at least 104-fold purification is required to isolate receptors. Solubilization with digitonin (Laduron and Ilien, 1982) and subsequent affinity chromatography using 3-(20 -aminobenzhydryloxy)tropane (Haga and Haga, 1983, 1985a) enabled purification of the muscarinic receptor. Reconstitution of the purified muscarinic receptor and the purified G protein Gi in phospholipid vesicles and demonstration of agonist-dependent activation of GTPase activity provided evidence that the muscarinic receptor interacts directly with and activates G protein Gi (Haga et al., 1985b, 1986). Partial proteolysis of the purified receptor, determination of partial amino acid sequences, and chemical synthesis of oligonucleotide probes corresponding to these amino acid sequences were used to identify the cDNA encoding the muscarinic receptor (Kubo et al., 1986a). Two types of cDNA were identified by this procedure: one was identified as the M1 subtype using M1 subtype-specific muscarinic ligands in conjunction with the tissue localization of the mRNA, while the other was cloned from porcine atria and identified as the M2 subtype (Kubo et al., 1986b). The cloned muscarinic receptors were shown to be homologous to the b adrenergic receptor (Dixon et al., 1986) and rhodopsin (Nathans and Hogness, 1983) in that they have the predicted 7-transmembrane (TM) topology. In addition, the b adrenergic receptor and rhodopsin have been shown to interact with different types of G protein, Gs and Gt, respectively. Subsequently, cDNAs of other muscarinic subtypes were identified using probes with sequences highly conserved among the previously cloned receptor sequences (Bonner et al., 1987, 1988; Peralta et al., 1988). The cloning of other GPCR genes demonstrated that the protein-coding region in many of these genes is intronless. Historically, both upper and lower case letters have been used to refer to muscarinic receptor subtypes, but it is now recommended to use the upper case designations M1, M2, M3, M4, and M5.

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. Figure 23-1 Structural formulas of acetylcholine, choline esters, and natural alkaloids that activate muscarinic receptors, and those that antagonize the action of acetylcholine

1.2 Distribution No strictly subtype-specific muscarinic antagonists have yet been reported. Moreover, muscarinic receptor subtypes are generally coexpressed with at least one other muscarinic receptor (Levey, 1993; Watson and Arkinstall, 2002). This posed problems in mapping their individual distributions using specific ligands alone. However, molecular cloning of muscarinic receptors has made it possible to prepare subtype-specific antibodies and oligonucleotide probes. Thus, the distributions of muscarinic receptor subtypes in both central and peripheral tissues have been investigated by immunohistochemical and northern blot or in situ hybridization studies along with pharmacological studies (Dorje et al., 1991; Wall et al., 1991; Levey, 1993; Wei et al., 1994). In the central nervous system, the M1 receptor is abundantly expressed in all forebrain areas, including the cerebral cortex, hippocampus, and striatum, and accounts for 40–50% of the total muscarinic receptors (Levey et al., 1991; Wall et al., 1991; Wei et al., 1994). Its distribution overlaps those of the M3 and M4 receptors. The M2 receptor accounts for over 80% of the total muscarinic receptors in the brainstem (Levey et al., 1991). The M4 receptor is expressed predominantly in the striatum (Bernard et al., 1992; Yasuda et al., 1993; Ince et al., 1997). The M5 receptor is expressed in several brain regions, such as the substantia nigra and ventral tegmental area, but it accounts for less than 2% of the total muscarinic receptors (Vilaro et al., 1990; Weiner et al., 1990; Yasuda et al., 1993). In the periphery, the M1 receptor is expressed in autonomic ganglia and certain secretory glands, such as the gastric glands. The M2 receptor is expressed mainly in the atrium. M2 receptors are also found in sympathetic ganglia, the ileum, uterus, bladder, and trachea. The M3 receptor is expressed in the pancreas, various smooth muscles, and secretory glands, such as the submaxillary gland. The M4 receptor is found in the lung (Dorje et al., 1991). The M5 receptor has been shown to be expressed in blood lymphocytes

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(Costa et al., 1995; Tayebati et al., 1999), skin fibroblasts (Buchli et al., 1999), and smooth muscles of the iris sphincter (Gil et al., 1997) and esophagus (Preiksaitis et al., 2000). The M5 receptor has also been identified in various peripheral and cerebral vessels (Phillips et al., 1997; Elhusseiny et al., 1999).

1.3 Diseases and Therapeutic Agents Related to Muscarinic Receptors Cholinergic neurons are lost selectively in Alzheimer’s disease, causing cognitive deficits. Systemic administration of a cholinesterase inhibitor has been reported to improve cognitive function, presumably by elevating acetylcholine levels in the brain (Standaert and Young, 2001; Terry and Buccafusco, 2003). Antimuscarinic agents are used for treatment of several diseases, including Parkinson’s disease (Standaert and Young, 2001), bronchial asthma, peptic ulcer, and overactive bladder (Brown and Taylor, 2001). The clinical use of muscarinic antagonists, however, is often limited by the frequent occurrence of unpleasant side effects caused by the low subtype-specificity of the ligands. For example, muscarinic antagonists, such as anti-pollakiuria agents, which are widely used in the elderly, cause dry mouth. Muscarinic agonists are used clinically in the treatment of glaucoma. More minor uses include suppression of atrial tachycardia, stimulation of intestinal motility and bladder emptying (Brown and Taylor, 2001).

2

Structure and Function

2.1 Structural Features Like most GPCRs, muscarinic receptors have been shown to possess seven hydrophobic segments, glycosylation sites in the N-terminal region, a disulfide bond between the second and third extracellular loops, and a palmitoylation site in the C-terminal region (> Figure 23-2) (Kubo et al., 1986a; Kurtenbach et al., 1990; Ohara et al., 1990; Savarese et al., 1992; Hayashi and Haga, 1997; Zeng et al., 1999a). X-ray crystallographic analysis of bovine rhodopsin in the ground state provided the first three-dimensional atomic-resolution structure of a GPCR and demonstrated that it is indeed composed of seven TM a-helices (TM1–TM7), connected by alternating intra- and extracellular loops, as well as a newly discovered 8th helix (H8) that projects from the end of TM7 and lies along the cytosolic surface of the cell membrane (Palczewski et al., 2000). The muscarinic receptors are also assumed to have seven TM a-helices, as was shown for rhodopsin, although direct evidence is still lacking: three-dimensional models of the M1 (Lu et al., 2001), M2 (Furukawa et al., 2002), and M3 (Li et al., 2005) receptors have been built via homology modeling using the atomic structure of bovine rhodopsin as a template (> Figure 23-3). One of the structural characteristics of muscarinic receptors is that each subtype contains a long third intracellular loop (i3) of 160–240 residues. Studies based largely on mutagenesis have shown that the Nand C-terminal portions of i3 play key roles in determining the specificity of the coupling between muscarinic receptors and G proteins (Wess et al., 1997) as described in > Sections 2.3 and 2.4). Most of the central part of i3 can be deleted without impairing the ability of the M2 receptor to couple with G proteins (Kameyama et al., 1994), indicating that only the N- and C-terminal portions of i3 are required for the interaction. One of the common functions of the long i3 is presentation of sites for agonist-dependent phosphorylation, as described in > Section 3. Structural characterization of M2i3 using circular dichroism and nuclear magnetic resonance (NMR) spectra suggests that the central part of i3 of the M2 receptor has a flexible structure (Ichiyama et al., 2006a). However, the possibility remains that i3 may form a definite structure when it binds cytosolic protein(s).

2.2 Agonist Binding All muscarinic receptor subtypes possess a conserved Asp residue in TM3 (Asp147 in the case of the M3 receptor; > Figures 23-2 and > 23-4). This Asp is modified irreversibly with propylbenzilylcholine mustard, an alkylating agent with a muscarinic antagonist function (Curtis et al., 1989). Mutation of this Asp results

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. Figure 23-2 Schematic diagram of the muscarinic receptor. Putative topology of the rat M3 receptor based on the topology prediction program, TMHMM (Krogh et al., 2001), the previous review (Wess et al., 1995), and homology modeling (Li et al., 2005). Glycosylation sites, a disulfide bond, and a palmitoylation site are also shown (Wess et al., 1995; Behr et al., 1998; Zeng et al., 1999a). The region in the C-terminal tail indicated with the shaded line is the putative 8th helix (H8). The shaded regions of the i2 and i3 loops are thought to contribute to agonist-dependent G-protein activation. Amino acid residues marked by open circles are conserved among most GPCRs. Residues highlighted with squares and the white letters on a black background are thought to be essential for G protein coupling and acetylcholine binding, respectively. The numbers given below the marked amino acids indicate their position in the rat M3 sequence. Only the membrane-proximal portions of the N-terminal domain, the i3 loop, and the C-terminal tail are shown. The omitted region in i3 in this figure corresponds to the central part of M2i3 that can be deleted without impairing the ability of the M2 receptor to couple with G proteins (Kameyama et al., 1994)

in loss of ligand-binding activity (Fraser et al., 1989). This Asp is conserved among amine receptors, such as adrenergic, dopamine, and serotonin receptors, and is thought to form an ionic bond with the positively charged head group of amine molecules (> Figure 23-4). Several site-directed mutagenesis studies have also identified other residues that contribute to agonist binding, which are found in the extracellular half of TM3–TM7 (> Figure 23-2) (Wess et al., 1991; Wess et al., 1992; Baldwin, 1993; Wess et al., 1993; Baldwin, 1994; Lu and Hulme, 1999; Lu and Hulme, 2000; Lu et al., 2001). In the M3 model structure, some residues, such as Asp147, Tyr148, Tyr506, and Tyr533, reside consistently in the vicinity of acetylcholine. In contrast, others, especially in TM4 and TM5, appear to lie far from acetylcholine, although they have been reported to be crucial for agonist binding (> Figure 23-4). These residues may contribute indirectly to agonist binding by affecting the local structure of the receptor or may be involved directly in agonist binding with the conformational change of the receptor.

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. Figure 23-3 Three-dimensional model of the muscarinic receptor. Stereo ribbon diagram of the rat M3 model structure drawn using the graphic program, PyMOL (DeLano, 2002), using the coordinates (PDB code, 2amk) built by homology modeling based on the X-ray structure of bovine rhodopsin (Li et al., 2005). Amino acid regions and colors are as follows: TM1 (amino acids 67–84), TM2 (104–124), TM3 (142–163), i2 (164–183), TM4 (184–206), TM5 (229–251), i3 (252–491), TM6 (492–512), TM7 (527–546), and H8 (548–558). The central part of i3 (263–477) is deleted in this model. Acetylcholine is drawn in space-filling mode in salmon pink (details of ligand-binding domain are given in the legend of > Figure 23‐4)

Some residues shown in > Figures 23-2 and > 23-4 are also required for antagonist binding, indicating that binding sites for antagonists overlap at least partially with those for agonists (Wess et al., 1991; Baldwin, 1993; Lu et al., 2001). Many agonists and antagonists for muscarinic receptors have been found from natural sources or synthesized artificially, but most show only low subtype specificity, reflecting the high degree of amino acid sequence homology around the agonist-binding sites among the five subtypes. Allosteric ligands that bind to regions other than the agonist-binding sites have also been developed (Ellis et al., 1993; Leppik et al., 1994; Matsui et al., 1995; Proska and Tucek, 1995; Birdsall et al., 1999). The binding of allosteric ligands is expected to be subtype-specific because relatively diverse amino acid sequences among the muscarinic subtypes contribute to their recognition (Tucek and Proska, 1995).

2.3 Coupling to G Proteins GPCRs are activated by binding of extracellular agonists, and the activated GPCRs interact with heterotrimeric G proteins causing a decrease in the affinity for GDP of the G protein a subunit (Ga). Dissociation of GDP is followed by binding of GTP to the Ga, which results in dissociation of the G protein into GaGTP and Gbg subunits (Roberts and Waelbroeck, 2004). GaGTP and Gbg each interact with effectors, such as

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. Figure 23-4 Ligand-binding interactions of the muscarinic receptor. The extracellular halves of TM helices of the rat M3 receptor are shown in a stereo ribbon diagram. It is bound with acetylcholine and is viewed from the intracellular side. The simulation model of acetylcholine was docked and refined using the program, DS-Modeling-SBD (Accelrys, San Diego, CA; Venkatachalam et al., 2003), with the assumption that the receptor is in the ground state and the docked acetylcholine takes the gauche configuration (see > Section 2.4). Residues that have been reported to be crucial for acetylcholine binding on the basis of mutagenesis studies and are highlighted in > Figure 23‐2 are drawn in stick mode. For simplicity, only the selected residues are numbered

adenylyl cyclase, phospholipase Cb (PLCb), and ion channels. G proteins are generally divided into four groups, Gs, Gi, Gq, and G12, depending on the species of Ga. The M1, M3, and M5 subtypes couple to Gq/11 type G proteins (Smrcka et al., 1991; Berstein et al., 1992a; Offermanns et al., 1994; Nakamura et al., 1995), which activate PLC, whereas the M2 and M4 subtypes couple to Gi/o type G proteins (Ikegaya et al., 1990; Parker et al., 1991; Offermanns et al., 1994), which are known to inhibit adenylyl cyclase activity. The molecular mechanisms by which individual receptors can selectively couple to a distinct subset of structurally similar G proteins have been investigated for many GPCRs. The region in the receptor that determines the G protein specificity appears to vary among GPCRs. In muscarinic receptors, studies with chimeric M2/M3 receptors have demonstrated that the N-terminal portion of i3 is critical in determination of the specificity of G protein coupling (> Figure 23-2) (Wess et al., 1989; Lechleiter et al., 1990; Wess et al., 1990; Wong et al., 1990). Based on a more systematic analysis of a large number of hybrid receptors, the second intracellular loop (i2) and the C-terminal segment of i3 have also been shown to contribute to the recognition of specific G proteins (> Figure 23-2) (Kunkel and Peralta, 1993; Moro et al., 1993; Wess et al., 1993; Blu¨ml et al., 1994a, b; Blin et al., 1995). Mutagenesis studies have identified some conserved residues essential for ligand-induced receptor activation, e.g., the DRY motif located at the N-terminus of i2 (Fraser et al., 1989; Zhu et al., 1994), Ile and Tyr residues located at the N-terminus of i3 (Blu¨ml et al., 1994a, b; Ho¨gger et al., 1995), and Lys residues located at the C-terminus of i3 (Ho¨gger et al., 1995; Lee et al., 1996) (> Figures 23-2 and > 23-5). In particular, several critical residues at the junction between i3 and TM6 of the M2 receptor (Val385, Thr386, Ile389, and Leu390, which correspond to Ala488, Ala489, Leu492, and Ser493, respectively, in the M3 sequence; > Figure 23-2) have been reported to recognize the C-terminus of Gai/o (Liu et al., 1995; Kostenis et al., 1997). These residues are involved in determining coupling selectivity and triggering G protein activation. There is ample evidence from biochemical and biophysical studies of muscarinic and other GPCRs that the N- and C-terminal segments of i3 form a-helical extensions of TM5 and TM6, respectively (Blu¨ml et al., 1994c; Altenbach et al., 1996; Hill-Eubanks et al., 1996; Ichiyama et al., 2006a). X-ray structural analysis of

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. Figure 23-5 Regions critical for G-protein recognition. The intracellular halves of TM helices of the rat M3 receptor are shown in a stereo ribbon diagram viewed from the intracellular side. Residues that have been reported to be crucial for G-protein recognition and are marked with squares in > Figure 23‐2 are drawn in stick mode. For simplicity, only the selected residues are numbered

rhodopsin verified the b-turn structure at the N-terminal end of i3, as suggested by NMR analysis of the synthetic i3 peptide of rhodopsin (Yeagle et al., 1995, 1997). In the M3 model shown in > Figure 23-3, the structure of the N-terminal part of i3 is assumed to be a b-turn following the a-helical extension of TM5.

2.4 Conformational Change upon Activation The three-dimensional structure of GPCR in an active state has not been determined. Spin labeling studies of rhodopsin have shown that when it is activated by light, TM6 rotates clockwise, as viewed from the cytoplasmic surface, and moves significantly outward relative to TM3 (> Figure 23-6) (Farahbakhsh et al., 1995; Altenbach et al., 1996; Farrens et al., 1996). Experiments involving introduction of fluorophores into b adrenergic receptors also suggest that TM6 moves outward when activated (Jensen et al., 2001; Lu et al., 2002). Moreover, fluorescence spectroscopic studies have shown that b2 adrenergic receptor activation also involves a conformational change that brings the cytoplasmic ends of TM6 and TM5 closer together (> Figure 23-6) (Ghanouni et al., 2001). In accordance with these findings, cross-linking of the cytoplasmic ends of TM3 and TM6 prevents receptor activation of these GPCRs (Sheikh et al., 1996, 1999). With regard to muscarinic receptors, it has been shown that activation of the M3 receptor leads to structural changes that cause the cytoplasmic ends of TM5 and TM6 to move closer together based on the in situ disulfide cross-linking strategy (> Figure 23-6) (Ward et al., 2002). Agonist-dependent rotational and/or translational movement of TM6 may cause exposure of the functionally critical residues at the juxtamembrane regions of i3 for interaction with the C-terminus of Ga. The interaction of a receptor and a G protein can be assessed as an agonist-dependent decrease in the affinity for GDP or guanine nucleotide-sensitive high-affinity agonist binding. The results of reconstitution experiments of the purified receptor and G proteins revealed that agonists bind to the muscarinic receptor with low affinity when G proteins are absent or GDP/GTP-bound G proteins are present, and that agonists bind to the receptor with high affinity when GDP/GTP-free G proteins are present (Florio and Sternweis, 1985; Haga et al., 1985b, 1986; Shiozaki and Haga, 1992). Recently, fusion proteins of GPCR with Ga have been used to characterize coupling between GPCR and G proteins. The GPCR-Ga fusion protein was first introduced and characterized for b2 adrenergic receptor and Gas (Bertin et al., 1994), followed by a number of fusion proteins between various types of GPCR and Ga protein (Seifert et al., 1999; Milligan, 2002).

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. Figure 23-6 Agonist-induced movement of TM6. The intracellular halves of TM helices of the rat M3 receptor are shown in cylindrical mode, viewed from the intracellular side. The results derived from spin labeling, fluorescence spectroscopic, and cross-linking studies can be explained by clockwise rotation of TM6 toward TM5 and/or its translational movement away from TM3 during agonist-induced activation of the receptor

The fusion proteins have a technical advantage in that the interaction between the receptor and Ga can be characterized easily using membrane preparations without purifying these proteins. The muscarinic receptor-Ga fusion protein can be functionally expressed in insect cells (Guo et al., 2001), mammalian cells (Suga et al., 2004), and in Escherichia coli (Ichiyama et al., 2006b). The M2-Gai1 fusion protein shows low, medium, and high affinities for GDP in the presence of agonist, partial agonist, and antagonist, respectively (Zhang et al., 2004). These results indicate that the function of ligands could be characterized by the affinity for GDP of Ga, with which the ligand-bound receptor interacts and activates. Attempts have been made to determine the active conformation of the muscarinic agonist using conformationally rigid acetylcholine analogs (Portoghese, 1970; Casy, 1975). In the majority of cases, the trans isomers of rigid analogs were more potent than the cis isomers in inducing muscarinic activities. Therefore, the pharmacologically active conformation of acetylcholine (> Figure 23-7Aa) and other analogs was assumed to take the trans O-C2-C1-N dihedral angle (180 ) (> Figure 23-7Bc) (Taylor and Insel, 1990). The conformations of two acetylcholine analogs, (S)-methacholine (> Figure 23-7Ab) and (2S,4R,5S)-muscarine (> Figure 23-7Ac), bound to the M2 receptor have recently been determined using NMR-TRNOE (transferred nuclear Overhauser effect). Contrary to expectations, the bound ligands were found to take the gauche O-C2-C1-N dihedral angle of þ60 (> Figure 23-7Bb) (Furukawa et al., 2002), which was close to those determined for the respective analogs in solution (þ80–85 ; > Figure 23-7Ba) (Casy et al., 1971; Furukawa et al., 2002) or for acetylcholine in a crystal (þ60 ; > Figure 23-7Bb) (Canepa et al., 1966). One possible explanation for the apparent discrepancy is that conformational change of the receptor-bound agonist may depend on the states of the receptor. NMR measurement was carried out in the absence of G proteins, and then the agonists should have bound to the receptor with low affinity, as described previously. It is possible that the agonist binds to the receptor with low affinity in the gauche conformation (þ60 ), whereas it takes the trans conformation (180 ) when bound to the receptor–G protein complex with high affinity in the absence of guanine nucleotides. Thus, we assume that the conformation

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. Figure 23-7 Structure and configuration of acetylcholine and its analogs. A, Structures of acetylcholine and its analogs. B, Configurations of (S)-methacholine in solution (a), bound to the M2 receptor without G protein (b), and predicted from the experiments using rigid analogs (c)

of the agonist bound to the receptor changes from the gauche to the trans form, thereby causing the conformational change of the receptor, which is accompanied by coupling of the agonist-bound receptor with G proteins and the concomitant release of GDP. In this assumption, the conformational changes of the agonist and the receptor are assumed to occur in a concerted manner together with the interaction of the receptor with G proteins, and the agonist-receptor-guanine nucleotide free-G protein complex is assumed to be a transition complex for the receptor-G interaction. The structure of the complex remains to be determined. GPCR-Ga fusion protein may be a target for purification and crystallization.

2.5 Regulation of Effector Proteins 2.5.1 Activation of Phospholipase C It has been shown that PLCb1 and PLCb3 are activated via the M1, M3, and M5 receptors and Gq in mammalian cells in culture. Reconstitution of the purified M1 receptor, purified Gq/11, and purified PLCb1 in lipid vesicles provided evidence that these three components are necessary and sufficient for acetylcholinedependent inositol-1,4,5-trisphosphate (IP3) production (Berstein et al., 1992b; Nakamura et al., 1995). At the same time, PLCb1 was found to stimulate hydrolysis of Gq/11-bound GTP acting as a GAP (GTPase-activating protein) of Gq/11 (Berstein et al., 1992b). PLCb2 is activated by the M2 receptor, probably through Gbg subunits released from Gi (Camps et al., 1992; Katz et al., 1992). PLCb3 is also activated by Gbg subunits (Carozzi et al., 1993; Murthy et al., 1996; Park et al., 1993).

2.5.2 Regulation of Adenylyl Cyclase Activation of the M2 and M4 receptors generally leads to inhibition of adenylyl cyclase (AC) through Gi/o, as described in > Section 2.3, although there have been several reports that muscarinic receptor activation leads to enhancement of AC, as described subsequently. To date, eight adenylyl cyclase isoforms have been

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identified, each of which is under diverse controls of activation and inhibition by G protein subunits, several kinases, and calmodulin (Gomperts et al., 2003). AC I is inhibited by Gai, Gao, Gaz, and Gbg, and AC VI is inhibited by Gai and Gaz (Taussig and Gilman, 1995). In the rat olfactory bulb, muscarinic receptors enhance AC activity by activating a pertussis toxin-sensitive G protein (Olianas and Onali, 1992). As AC II, which is expressed abundantly in the olfactory bulb, is activated synergistically by Gas and Gbg, AC activation in the olfactory bulb may be mediated by Gbg released from Gi/o by the action of M2 and/or M4 receptors. In human neuroblastoma cells, Ca2+ and protein kinase C (PKC) are required for increases in cAMP levels mediated by M1 and M3 receptors (Jansson et al., 1991; Baumgold et al., 1992), probably through activation of AC I and AC III, both of which can be activated by Ca2+/calmodulin.

2.5.3 Regulation of Ion Channels The G protein-coupled, inwardly rectifying K+ (GIRK) channel is activated by the M2 receptor. In this case, Gbg subunits released from Gi bind to a region on the cytoplasmic domain of the channel, causing it to open (Logothetis et al., 1987; Kofuji et al., 1995). One of the voltage-sensitive K+ currents is the M current, which is found in the nervous system, such as the sympathetic ganglion and pyramidal cells of the hippocampus. The M current was so termed because it is inhibited by acetylcholine via muscarinic receptors, although it is also blocked by other transmitters acting on GPCRs that are associated with PLC (Brown and Adams, 1980). The KCNQ2/KCNQ3 heteromeric M channel plays the central role in producing the M current (Wang et al., 1998). Recently, the cAMPdependent protein kinase (PKA) anchoring protein, AKAP79/150, has been shown to function as a scaffolding protein to form a complex with the M1 receptor, PKC, and M channel to direct muscarinic activation efficiently toward suppression of the M current through phosphorylation of the KCNQ2 subunit by PKC (Hoshi et al., 2005). It has been shown that pertussis toxin-sensitive G proteins inhibit N-type Ca2+ channels in a manner independent of soluble intracellular messengers. Gbg subunits also directly modulate Ca2+ channels in sympathetic neurons (Herlitze et al., 1996; Ikeda, 1996). Meanwhile, other Ca2+ channels can be opened indirectly through activation of PLC. The Ca2+ increase caused by muscarinic receptors consists of two phases: a transient spike-shaped increase, and a subsequent long-lasting plateau. The first rapid increase is produced by release of Ca2+ from IP3-sensitive intracellular Ca2+ stores, and the subsequent plateau is maintained by Ca2+ influx from the extracellular space through Ca2+ channels on the plasma membrane (Clapham, 1995a, b; Zhang and Saffen, 2001; Bolotina and Csutora, 2005; Ebihara et al., 2006).

3

Regulation of Receptor Activity

3.1 Desensitization Prolonged exposure of receptors to agonists results in diminished responsiveness, which is referred to as receptor desensitization. The process in which only agonist-bound receptors are desensitized is called homologous desensitization, while the process in which not only the agonist-bound but also agonist-free receptors are desensitized is called heterologous desensitization. Receptor desensitization includes three processes: uncoupling (decrease in coupling efficiency to G proteins), receptor internalization, and downregulation (degradation of the receptor). These processes are initiated in many cases by phosphorylation of the receptor catalyzed by PKA, PKC, and G protein-coupled receptor kinase (GRK). PKA and PKC phosphorylate both agonist-bound and agonist-free receptors (Haga et al., 1988, 1990a, 1993; Uchiyama et al., 1990). In contrast, GRK phosphorylates only the agonist-bound receptors (Haga and Haga, 1989; Haga et al., 1994a). The former two kinases are thought to contribute to heterologous desensitization, whereas the latter contributes to homologous desensitization.

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3.2 Phosphorylation by GRK and Receptor Internalization Seven GRK subtypes, GRK1–GRK7, have been identified (Penn et al., 2000). In most cases, GRK fulfills its function through the agonist-dependent phosphorylation of GPCRs. For example, the M2 receptor is phosphorylated in an agonist-dependent manner by GRK2 (Richardson and Hosey, 1992; Kameyama et al., 1993; Haga et al., 2002), GRK3 (Richardson et al., 1993), and GRK5 (Kunapuli et al., 1994). The M3 receptor is also phosphorylated by GRK2 in an agonist-dependent and Gbg-enhanced manner; however, neither GRK5 nor GRK6 significantly phosphorylates the M3 receptor (Debburman et al., 1995). GRK2 shows a marked preference for negatively charged amino acids localized at the N-terminal side of the target Ser or Thr residue of the receptor (Onorato et al., 1991). It is less likely that GRK2 discriminates the rigid local tertiary structure of the target sequence, because i3 is likely to be a flexible loop with little secondary structure (Ichiyama et al., 2006a). GRK2 binds to Gbg subunits via its PH (pleckstrin homology) domain located in the C-terminus, thereby translocating itself from the cytosol to the inner surface of the plasma membrane and being activated synergistically by the agonist-bound receptor and Gbg (Haga and Haga, 1990b, 1992, 1994b; Pitcher et al., 1992; Kameyama et al., 1993; Touhara et al., 1994). The Gbg-binding site in GRK2 has been determined by X-ray crystallography. The structure of GRK2 in complex with Gbg subunits shows how RH (regulator of G protein signaling homology), PH, and kinase domains arrange themselves at the cell membrane (Lodowski et al., 2003). The fact that phosphorylation of a receptor by GRK2 promotes internalization of the receptor was first reported for the M2 receptor (Tsuga et al., 1994, 1998a, b, c). Sites for phosphorylation of muscarinic receptors by GRK2 exist in i3 (Nakata et al., 1994). It is generally assumed that, once a receptor is phosphorylated, arrestin binds to the phosphorylation site(s) with high affinity, and subsequently exerts its effects in the desensitization processes in two ways. First, arrestin prevents the receptor from coupling to G proteins. Second, it can serve as an adaptor protein to link the receptor to clathrin-coated pits, inducing receptor internalization. Internalized receptors may be degraded, probably in lysosomes (down-regulation), or may return to the cell surface (receptor recycling). M2 and M4 receptors show different recycling patterns. While approximately 60% of internalized M4 receptors are recycled back to the cell surface following removal of agonist, at most 30% of internalized M2 receptors are recycled. When the chimeric receptors in which i3 is exchanged between the two subtypes are subjected to internalization and recycling, the mutant M2 receptor having M4i3 instead of M2i3 shows the ‘‘M4-type’’ recycling pattern (Hashimoto et al., 2006), suggesting that i3 plays a critical role in recycling of muscarinic receptors.

3.3 Regulation by RGS Proteins GTP bound to Ga is hydrolyzed by the intrinsic GTPase activity, which determines the lifetime of the active state of a G protein. The hydrolysis of GTP bound to Ga in vivo is accelerated by binding of G proteins to their effector proteins, e.g., PLCb for Gq and p115RhoGEF for G13, or to a growing family of RGS proteins (regulators of G protein signaling). These proteins show GAP activity for heterotrimeric G proteins (Ross and Wilkie, 2000). GAP activity facilitates the termination of a signal upon removal of the stimulus. There are over 30 RGS proteins, all of which have an RGS domain consisting of 130 amino acids. The atomic structure of RGS4 forming a complex with Gai1-Mg2+ -GDP-AlF4 has been determined (Tesmer et al., 1997). RGS4 appears to catalyze rapid hydrolysis of GTP primarily by stabilizing the transition state for GTP hydrolysis. GRKs have an RH domain at their N-terminus. GRK2 binds to the activated Gaq through its RH domain and may regulate receptor–G protein interaction in a phosphorylation-independent manner (Dhami et al., 2004; Tesmer et al., 2005). RGS proteins can function as scaffold proteins for complex formation with various proteins, including Ga and receptors at the cytosolic side of the membrane (Hepler, 2003). RGS2 and RGS4 bind directly to M1i3 through its N-terminal region to attenuate Gq signaling (Bernstein et al., 2004). Very recently, RGS8, but not its splice variant RGS8S with a different N-terminus, has also been shown to bind directly to M1i3 via its N-terminus to suppress Gq signaling (Itoh et al., 2006).

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It remains to be clarified whether RGS2, RGS4, and RGS8 have different affinities for M1i3 or any functional differences in regulating the M1 receptor function. It is tempting to speculate that the cytosolic surface of GPCRs may selectively recruit specific RGS protein(s) to direct RGS activity toward the specific GPCR.

4

Dimerization/Oligomerization

The possibility of dimerization/oligomerization for class-A (rhodopsin-family) GPCRs, and its possible effects on ligand-binding properties, G-protein signaling, membrane translocation, and intracellular trafficking have been discussed (Milligan, 2004; Nakata et al., 2004). Dimerization/oligomerization of GPCRs have been examined by various techniques, such as dominant-negative effect of the inactive mutant receptor on the wild-type molecule, co-transport of two different receptors by activation with one specific agonist, co-immunoprecipitation of two differentially tagged receptors with one specific antibody, FRET (fluorescence resonance energy transfer) measurements, etc. From atomic-force microscopy studies, it seems that rhodopsin forms arrays of dimers in native disk membranes. Thus, reports that class-A receptors may form dimers and/or higher-order oligomers appear to be growing. Meanwhile, there are indications that dimerization/oligomerization of GPCRs can occur as artifacts due to overexpression of GPCRs on cultured cells (Salim et al., 2002; Javitch, 2004). The physiological significance of dimerization/ oligomerization remains to be determined. Experiments using M3 and a2C chimeric receptors suggested that the muscarinic receptor may form dimers (Maggio et al., 1993). Two types of chimera, M3(TM1–5)-a2C(TM6,7) and a2C(TM1–5)M3(TM6,7), in which a region containing TM6 and TM7 was exchanged between the two receptors, were expressed together in cultured cells. Significant numbers of muscarinic and adrenergic binding sites were detected after coexpression of the two chimeric receptors, whereas neither of the mutant receptors showed ligand-binding activity when expressed alone. These sites showed ligand-binding properties similar to those of the respective wild-type receptors. Carbamylcholine (muscarinic agonist, > Figure 23-1) stimulation of cells cotransfected with the two hybrid receptors caused significant hydrolysis of phosphatidyl inositol, although the maximum amount of inositol phosphates produced was approximately half of that produced by the wild-type M3 receptor, whereas stimulation of cells transfected with either of the mutant receptors did not. The EC50 value determined for the cotransfected chimeric receptors (4.3 mM) was only 3-fold higher than that for the wild-type M3 receptor (1.5 mM). These results suggest that the two chimeric proteins interact with each other and form the respective ligand-binding and G-protein recognition sites, i.e., functional complementation. Furthermore, Wess et al. reported that M3 receptors solubilized with digitonin from COS-7 cells exist as a mixture of monomers and dimers, as demonstrated by SDSpolyacrylamide gel electrophoresis in the absence of reducing agents, and convert entirely to the monomeric form in the presence of dithiothreitol (Zeng and Wess, 1999b). They suggested that Cys140 and Cys220 play crucial roles in the formation of M3 dimers. The functional differences, if any, in muscarinic receptor monomers and dimers remain to be determined.

5

Complex Formation with Other Proteins

Recently, agonist-dependent transient complex formation involving the M1 receptor, TRPC6 (transient receptor potential-canonical subtype 6) channel, PKC, immunophilin FKBP12, and calcineurin/calmodulin has been reported (Kim and Saffen, 2005). This study, based on immunoprecipitation of endogenous proteins, provided new insight into the molecular rearrangements and phosphorylation/dephosphorylation of TRPC6 channels following GPCR activation. The formation of a supramolecular complex is essential in that the highly ordered proteins contained in the complex interact with each other and transmit signals with high accuracy.

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Knockout Studies

A series of knockout (KO) mice deficient in one or more of five subtypes have recently been generated and characterized to reveal the functions of individual subtypes as well as the involvement of the receptor in specific diseases (Bymaster et al., 2003; Wess et al., 2003; Matsui et al., 2004; Wess, 2004). Mutant mice lacking each of the muscarinic receptors are viable, fertile, and appear generally healthy, although detailed physiological, behavioral, and biochemical studies have revealed the characteristic phenotypic deficits or changes in these KO mice as described later. Disruption of one specific muscarinic subtype gene does not seem to have major effects on the expression levels of the remaining four muscarinic subtypes (Hamilton et al., 1997; Gomeza et al., 1999a, b; Miyakawa et al., 2001; Yamada et al., 2001a), suggesting that compensatory changes in the expression levels of muscarinic receptors are less likely to have a major impact on the outcome of mouse phenotyping studies.

6.1 M1 KO Mice M1 KO mice are resistant to the seizures evoked by systemic administration of the muscarinic agonist pilocarpine (> Figure 23-1; Hamilton et al., 1997). M1 KO mice also show significantly increased locomotor activity, probably due to the elevated dopaminergic transmission in the striatum (Gerber et al., 2001). This seems consistent with the observation that M1-preferring antagonists are useful in treatment of Parkinson’s disease, a brain disorder characterized by reduced striatal dopamine levels (Gerber et al., 2001; Miyakawa et al., 2001). As the muscarinic antagonist scopolamine (> Figure 23-1) causes amnesia and Alzheimer’s disease is associated with impairment of cholinergic neurons (Bartus et al., 1982; Coyle et al., 1983), it is of particular interest whether mutant mice lacking the M1 receptor, which is expressed abundantly in forebrain areas as described in > Section 1.2, show impaired learning and memory. However, M1 KO mice appear to exhibit only moderate cognitive impairments (Miyakawa et al., 2001; Anagnostaras et al., 2003). Thus, it remains to be clarified whether muscarinic receptors are critical in learning and memory.

6.2 M2 KO Mice M2 KO mice are deficient in agonist-dependent akinesia and tremor, implying that striatal M2 receptors are involved in Parkinson’s disease (Gomeza et al., 1999a). The agonist-induced hypothermic responses are significantly reduced in M2 KO mice, indicating that the M2 receptor participates in regulation of body temperature (Gomeza et al., 1999a). The agonist-mediated bradycardic responses are completely abolished in M2 KO mice, confirming that the M2 receptor is involved in attenuation of heartbeat. M2 KO mice also show reduced agonist-induced contractile responses in the stomach, urinary bladder, and trachea, indicating that the M2 receptor plays a role in agonist-dependent smooth muscle contractility (Stengel et al., 2000).

6.3 M3 KO Mice M3 KO mice show the most significant phenotypic differences from the wild-type (WT) mice as compared with other single KO strains. M3 KO mice are characterized by severe functional impairment of multiple organs innervated by parasympathetic nerves (Matsui et al., 2000). M3 KO mice show growth retardation (Matsui et al., 2000; Yamada et al., 2001a). Matsui et al. proposed that this growth retardation is caused mainly by insufficient food intake due to reduced salivation based on the observations that salivation induced by a moderate dose of pilocarpine is nearly abolished, wet paste feeding improves the growth of KO mice, and the mutant mice approach the water nozzle very frequently when they eat dry pellet food compared to WT controls (Nakamura et al., 2004). In contrast, Wess et al. hypothesized that the lean phenotype is probably caused by central anorexia resulting from hypothalamic

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dysfunction, such as reduced expression levels of melanin-concentrating hormone, an appetite-stimulating peptide (Yamada et al., 2001a). Thus, it remains unclear to what extent these peripheral and central deficits contribute to the reduced body weight and food intake displayed by M3 KO mice. M3 KO mice, especially males, show prominent distension of the urinary bladder. The M3 receptor plays the principal role in cholinergic contractility of detrusor (Matsui et al., 2000) and ileal smooth muscles (Matsui et al., 2002). However, M3 KO mice show no apparent signs of constipation or ileus. Although isolated preparations of ileal smooth muscles derived from M2/M3 KO mice completely lack cholinergic contraction, these double KO mice show no apparent intestinal complications (Matsui et al., 2002). The pupil size of M3 KO mice is significantly larger than that of WT controls (Matsui et al., 2000), while the pupil size of M2/M3 KO mice is smaller than that of M3 KO mice (Matsui et al., 2002). These observations indicate that the M2 and M3 receptors play roles in mydriasis and myosis, respectively, in regulation of pupil diameter.

6.4 M4 KO Mice The M4 receptor has been shown to contribute to inhibition of AC activity in the striatum, which is stimulated by dopamine receptors (Olianas et al., 1996). M4 KO mice consistently show a small but significant increase in basal locomotor activity and locomotor stimulation with D1 dopamine receptor agonists is greatly enhanced in M4 KO mice (Gomeza et al., 1999b), suggesting that striatal M4 receptors exert an inhibitory effect on D1 receptor-stimulated locomotor activity. Systemic administration of scopolamine suppresses the cataleptic response induced by haloperidol (D2type dopamine receptor antagonist) in WT mice. In contrast, scopolamine shows no such effect in M4 KO mice (Karasawa et al., 2003). This result supports the suggestion that motor dysfunction can be caused by an imbalance between dopaminergic and cholinergic actions.

6.5 M5 KO Mice In M5 KO mice, dopamine release in the nucleus accumbens, which is initiated by stimulation of the cholinergic neurons in the laterodorsal tegmental nucleus, is partially lost (Forster et al., 2002). Thus, the M5 receptor on midbrain dopamine neurons is thought to mediate facilitation of dopamine release in the nucleus accumbens. M5 KO mice also lack acetylcholine-mediated dilation of cerebral arteries and arterioles (Yamada et al., 2001b).

7

Conclusions

In the long history of physiological and biochemical studies on acetylcholine extending back to the nineteenth century, the acetylcholine receptor had been a functional concept rather than a molecular entity before the cDNAs and genes whose products are responsible for nicotinic and muscarinic actions were identified in 1980–1990. Molecular characterization of the related genes and proteins provided evidence that muscarinic receptors, G proteins, and effector proteins are distinct functional proteins. Furthermore, there are increasing examples of supramolecular complex formation of muscarinic receptors with various membrane and/or cytosolic proteins. These accessory proteins have been implicated in modulating signaling properties of the receptor, as well as in trafficking of the receptor to and from the plasma membrane. Thus, future studies should take into account that a receptor can be regulated in a spatiotemporal manner following multiprotein complex formation. With the development of structural biology, it has become possible to elucidate the tertiary structures of signaling proteins, leading to understanding of the signaling pathways across the plasma membrane on a structural basis. In particular, the atomic structure of rhodopsin has provided a solid basis for constructing

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a structural model of muscarinic receptors and other GPCRs, helping interpret large amounts of mutagenesis data. It is important to reveal the tertiary structure of muscarinic receptors to clarify agonist and antagonist binding modes. Development of ligands with high subtype specificity based on the structural information would be useful for the treatment of various diseases related to muscarinic receptors. A wealth of novel insights into the physiological roles of the individual muscarinic subtypes have been obtained by phenotypic analyses of M1–M5 KO mice. Development of KO mice lacking two or more muscarinic subtypes or those in which a specific muscarinic subtype is inactivated in a conditional manner would provide invaluable research tools. Information on the physiological function of each muscarinic subtype as well as its precise distribution would contribute to the development of novel muscarinic medications useful in a variety of pathophysiological conditions.

Acknowledgment We thank Dr. Hiroyuki Nakamura (Gakushuin Univ.) for his support in performing the docking simulation of acetylcholine into the M3 receptor model.

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Structure of IP3 Receptor

H. Yamazaki . K. Mikoshiba

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 442

2 2.1 2.2 2.2.1 2.2.2 2.2.3 2.2.4

IP3R Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 442 Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 442 Splice Variants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 443 SI in IP3R1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 443 SII in IP3R1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 446 SIII in IP3R1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 446 SIm2 and TIPR in IP3R2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 446

3 3.1 3.1.1 3.1.2 3.1.3 3.1.4 3.1.5 3.2 3.3

Domain Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 447 Five-Domain Structure Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 447 N-Terminal Coupling Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 447 Ligand Binding Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 447 Internal Coupling Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 450 Transmembrane Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451 Gatekeeper Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451 Trypsin Digestion Pattern Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 452 Head-To-Tail Interaction Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 452

4

Three-Dimensional Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 453

5

Functional Properties of IP3R Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 453

6

Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 455

#

2009 Springer ScienceþBusiness Media, LLC.

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Structure of IP3 receptor

Abstract: The inositol 1,4,5-trisphosphate receptor (IP3R) is an intracellular tetrameric calcium (Ca2+)release channel localized predominantly on the endoplasmic reticulum of all cell types. IP3R releases Ca2+ into the cytoplasm in response to inositol 1,4,5-trisphosphate (IP3) produced by diverse stimuli, generating complex local and global Ca2+ signals that regulate numerous cell physiological processes, including gene transcription, secretion, and synaptic plasticity (see Chapter 25). This versatility is derived from the diverse properties of IP3R. Three IP3R isoforms in mammals and a single isoform in invertebrates are expressed with several splice variants. Though they have conserved functional domains, IP3R channel activity is regulated in an isoform-specific manner by IP3 and other ligands, such as Ca2+ and ATP, and also by multiple interacting proteins. Recent biochemical, electrophysiological, and structural analyses have deepened our understanding of the functional properties of IP3R. Herein, we review the genetic and functional structures of IP3R and discuss the functional diversity of IP3R isoforms.

1

Introduction

It is well established that Ca2+ plays an important role in many physiological phenomena in a variety of cells. Through the late 1980s, knowledge accumulated that a second messenger, IP3, binds to unidentified ‘‘IP3 receptor’’ and induces Ca2+ mobilization from internal stores, mainly the endoplasmic reticulum (Berridge and Irvine, 1989). Remarkable IP3-binding activity was found in the cerebellum (Worley et al., 1987a, b), and IP3-binding protein was highly purified from the cerebellum (Supattapone et al., 1988). The purified protein was reconstituted into lipid vesicles, and it was clarified that the ‘‘IP3 receptor’’ itself is a Ca2+ release channel (Ferris et al., 1989). Independent of these IP3 signaling studies, IP3R protein was characterized by several groups as a membrane glyco- and phospho-protein, which is abundant in the cerebellum, and it was variously named P400 (Mallet et al., 1976; Mikoshiba et al., 1979, 1985), PCPP-260 (Walaas et al., 1986), or GP-A (Groswald and Kelly, 1984). P400 protein was found to be deficient in hereditary cerebellar ataxic mutant mice, specifically in degenerating Purkinje cells (e.g., Purkinje-cell-degeneration, staggerer, and nervous) (Mallet et al., 1976; Mikoshiba et al., 1979, 1985). Using purified P400 proteins, anti-P400 monoclonal antibodies were obtained (Maeda et al., 1988, 1989). By immunoscreening mouse cerebellum cDNA libraries with these antibodies, the cDNA of P400 (IP3R) was primarily cloned (Furuichi et al., 1989). The partial cDNA was also cloned through differential subtraction analysis of cerebellar cDNAs between wild-type and Purkinje-cell-degeneration mutant mice (Nordquist et al., 1988; Mignery et al., 1989). The cloned P400 cDNA was expressed in the mouse fibroblast L cell line, and the coded protein was confirmed to be IP3R from its IP3 binding and Ca2+ release activity (Miyawaki et al., 1990). Agarose-PAGE analysis of the cross-linked IP3R revealed the IP3R channel to be a tetramer composed of four non-covalently bound IP3R molecules (Maeda et al., 1991).

2

IP3R Family

IP3R has been found in organisms from nematodes to humans, and consists of three isoforms and various splice variants. The isoform-specific channel properties, modulations by interacting factors, and expression profiles generate the diversity of Ca2+-mobilization patterns in individual cells.

2.1 Isoforms From sequence homology, three IP3R isoforms (IP3R1, IP3R2, and IP3R3) encoded on distinct genes have been cloned in mammals (IP3R1: [mouse] Furuichi et al., 1989; [rat] Mignery et al., 1990; [human] Yamada et al., 1994, Harnick et al., 1995, and Nucifora et al., 1995. IP3R2: [mouse] Iwai et al., 2005; [rat] Su¨dhof et al., 1991; [human] Yamamoto-Hino et al., 1994. IP3R3: [mouse] Iwai et al., 2005; [rat]

Structure of IP3 receptor

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Blondel et al., 1993; [human] Maranto, 1994, and Yamamoto-Hino et al., 1994). In Xenopus laevis, only one IP3R isoform has been cloned among three isoforms (Kume et al., 1993). On the other hand, there is a single IP3R gene in Drosophila melanogaster (Yoshikawa et al., 1992; Hasan and Rosbash, 1992), the starfish Asterina pectinifera (Iwasaki et al., 2002), the lobster Panulirus argus (Munger et al., 2000), and Caenorhabditis elegans (Baylis et al., 1999). The IP3R family members share 40–70% amino acid sequence identity, except for the cloned Xenopus IP3R, which is highly homologous to mammalian IP3R1 sharing about 90% identity. Homologies among the three isoforms of mammalian IP3R are 60–70% identity, and the same isoforms of different species share more than 95% identity. IP3R has some sequence homology with another intracellular Ca2+ channel, ryanodine receptor protein, especially the N-terminal region and the membrane-spanning region (Furuichi et al., 1989; Henzi and MacDermott, 1992; Ponting, 2000).

2.2 Splice Variants Splice variants have been identified as SI, SII, and SIII in IP3R1, and SIm2 and TIPR in IP3R2. In IP3R1, 48 possible splice variants from at least 96 distinct transcripts have been estimated and, to date, 17 structural variations of IP3R1 have been detected in the rat brain (Regan et al., 2005). The splicing occurs unequally not only in various tissues but also in developmental stages of various brain parts (Nakagawa et al., 1991a, b; Regan et al., 2005). Invertebrates appear to express only a single IP3R isoform with several splice variants. The Caenorhabditis eleganse IP3R has three distinct amino-termini and two splice sites (Baylis et al., 1999; Gower et al., 2001) and the Drosophila melanogaster IP3R exists as two alternatively spliced forms (Sinha and Hasan, 1999).

2.2.1 SI in IP3R1 The SI region (15 amino acids, residues 318-VDPDFEEECLEFQPS-332 of the mouse IP3R1 (mIP3R1) sequence [Mignery et al., 1990, Nakagawa et al., 1991a, Harnick et al., 1995]) is located on the loop between the b6- and b7-strands of the ligand-binding domain (> Figures 24-1a and > 24-2c). The SI– form

. Figure 24-1 Structure model of IP3R. (a) Limited trypsin digestion pattern and domain structure of IP3R1 with associated molecules. Residues are numbered according to the mouse IP3R1 (SI+, SII+, SIII) sequence (NCBI accession number X15373). The molecular mass of major tryptic fragments (Yoshikawa et al., 1999) is indicated in kDa. Ligand binding domain and membrane-spanning regions (M1–6) are indicated by black boxes, and the channel pore position is indicated by a gray box. Alternative splicing sites SI, SII, and SIII of IP3R1 and the corresponding site of the splicing site SIm2 of IP3R2 are indicated. Ca, putative Ca2+ binding sites (Sienaert et al., 1996, 1997); P, phosphorylation sites (Y353 is a Fyn tyrosine kinase phosphorylation site, S421 and T799 are cdc2/CyB phosphorylation sites, S1588 and S1755 are PKA/PKG phosphorylation sites, and S2681 is an Akt phosphorylation site [Foskett et al., 2007]); two branched bars in the transmembrane domain, N-glycosylation sites N2475 and N2503 (Michikawa et al., 1994). ATPA, ATPB, and ATPC are ATP binding sites (Wagner et al., 2006). Possible binding sites of the interacting proteins are shown with arrows and gray horizontal lines (Patterson et al., 2004a; Foskett et al., 2007). (b) Transmembrane topology and gating model of IP3R. Two IP3R monomers of the tetramer channel are shown. The C (conserved) region of the M5–M6 loop contains a pore helix and selectivity filter sequences, which comprise a channel pore. In an open state, the bending of the M6 transmembrane helix at Gly2586, a residue which is conserved among IP3R and RyR families (Shah and Showdhamini, 2001), is estimated based on the analogy of a ‘‘gating hinge’’ of potassium channels (Jiang et al., 2002b; MacKinnon, 2003). The N-terminal coupling domain suppresses IP3-binding to the ligand binding domain at low IP3 concentrations under unstimulated conditions. With IP3-generating stimuli, IP3 overcomes this suppression and IP3-binding induces the conformational change in IP3R, that gates the channel opening through the headto-tail interactions among the N-terminal coupling domain, the M4–M5 linker, and the gatekeeper domain (see the online version at www.SpringerLink.com)

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. Figure 24-1 (continued)

increases during development of the cerebellum (Nakagawa et al., 1991a; Regan et al., 2005). The SI region exists within one of the putative Ca2+ binding sites of IP3R1 (residues 304–381 of mIP3R1) (Sienaert et al., 1997). Without Ca2+, no significant difference in IP3 affinity between the SI splice variants has been observed (Newton et al., 1994; Lievremont et al., 1996). However, at a 1-mM-free Ca2+ concentration, IP3-binding to the SI– form is inhibited markedly sharper than the SI + form, resulting in more oscillatory behavior of cytoplasmic Ca2+ in cells expressing IP3R1 (SI–) (Regan et al., 2005).

. Figure 24-2 Crystal structure of the N-terminal coupling and ligand binding domains of mouse IP3R1 (mIP3R1) and the region responsible for IP3-binding suppression. Ribbon diagram (a) and surface representation (b) of amino acid residues 2–223 of mIP3R1 (PDB code 1XZZ). The corresponding region of the SIm2 splicing site of mIP3R2 is colored in orange in a. Conserved amino acid residues among the IP3R family are plotted with a color gradient from magenta (identical residues) to green (least conserved residues) in b (adapted from Bosanac et al., 2005). Ribbon diagram (c) and surface representation (d) of amino acid residues 224–604 of mIP3R1 (PDB code 1N4K). SI splicing site and limited trypsin digestion site are indicated in c. Surface electrostatic potential with positive charge depicted in blue in d (adapted from Bosanac et al., 2002). The IP3 dissociation constants for wild-type T604, an IP3R N-terminal construct (residues 1–604) containing both the N-terminal coupling and ligand binding domains, and various site-directed mutants (e). *p < 0.05 and **p < 0.01 (Student’s t test, compared with the value of T604). Residues involved in IP3binding suppression determined in e and key sites for binding other proteins (Bosanac et al., 2005) are shown in f (see the online version at www.SpringerLink.com)

Structure of IP3 receptor

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2.2.2 SII in IP3R1 The SII region (40 amino acids, residues 1692–1731 of mIP3R1 [Danoff et al., 1991; Nakagawa et al., 1991a; Nucifora et al., 1995]) is located in the internal coupling domain (> Figure 24-1a), and is subdivided into three splicing subregions, A (23 amino acids, residues 1692-QISIDESENAELPQAPEAENSTE-1714), B (one amino acid, residue Q-1715), and C (16 amino acids, residues 1716-ELEPSPPLRQLEDHKR-1731). The A subregion of SII is also reportedly comprised of two variably spliced exons, SIIA1 (the former 11 amino acids) and SIIA2 (the latter 12 amino acids) (Regan et al., 2005). Neuronal IP3R1 has variable SII subtypes, e.g., SII+, SII, SIIB (only subregion B is deleted), and SIIBC– (subregions B and C are both deleted) (Nakagawa et al., 1991a, b). On the other hand, non-neuronal IP3R is mainly the SII– subtype and in some cell lines SIIA– was also reported (Iida and Bourguignon, 1994). The SII sequence is absent from the IP3R2 and IP3R3 isoforms. The SII sequence is located between two PKA/PKG phosphorylation consensus sites (Ser-1588 and Ser-1755) (Danoff et al., 1991; Ferris et al., 1991; Haug et al., 1999; Soulsby et al., 2004) and splicing affects the phosphorylation kinetics and these sites (Danoff et al., 1991; Ferris et al., 1991), consequently regulating the Ca2+ release activity of IP3R (Wagner et al., 2003, 2004). Interestingly, splicing out of the SII region creates an ATP-binding site (GxGxxG) (ATPC site in > Figure 24-1a), and bound ATP increases channel activity probably via activation of PKA-mediated phosphorylation of IP3R (Wagner et al., 2006). The SII region is also in the vicinity of a putative calmodulin (CaM) binding domain, residues 1564–1585 of mIP3R1 (SI+, SIII) (Yamada et al., 1995). The SII deletion causes stronger affinity to CaM, and binding is inhibited by phosphorylation with PKA (Lin et al., 2000). Furthermore, the splicing alters Ca2+ dependencies of IP3 binding and channel activity (Lin et al., 2000; Tu et al., 2002). The opisthotonos (opt) spontaneous mouse mutation resulting in epileptic-like behaviors, which are postnatally lethal, was identified as a deletion in the next SII segment, residues 1732–1839 of mIP3R1 (SI+, SII+, SIII) (Street et al., 1997). This deletion causes some changes in the Ca2+-release kinetics of IP3R, though the relationship between the opt mutation and SII splicing remains uncertain (Tu et al., 2002).

2.2.3 SIII in IP3R1 The SIII region (9 amino acids, residues 904-NNDVEKLKS-912 of human IP3R1 SI- sequence [Nucifora et al., 1995], corresponding to an insertion between the residues 917 and 918 of mIP3R1 SI+) is located in the internal coupling domain (> Figure 24-1a). The SIII+ form predominates in most brain regions, except the adult cerebellum, while the SIII – form predominates in peripheral tissues (Nucifora et al., 1995, Regan et al., 2005). Functional diversity in SIII splicing was proposed as a means of differential creation of protein kinase C (PKC)-binding sites by which the SIII+ acquires a new PKC potential site (KLKS) in the SIII insert but the SIII– has a potential site (NKGS) just at the joining point after deletion of the SIII sequence (Nucifora et al., 1995).

2.2.4 SIm2 and TIPR in IP3R2 The SIm2 region (33 amino acids, residue 176-IVVGDKVVLMPVNAGQPLHASNVELLDNPGCKE-208 of mIP3R2 [Iwai et al., 2005], corresponding to a deletion between residues 176 and 208 of mIP3R1 extends from the upstream region of the b9-strand to just before the b11-strand of the N-terminal coupling domain (> Figures 24-1a and > 24-2a). The SIm2– has neither IP3 binding activity nor Ca2+ release activity. The expression level of the SIm2– form is low in various tissues, though its coexpression may affect intracellular Ca2+ mobilization (Iwai et al., 2005). Another mouse IP3R2 splice variant termed TIPR was detected, in skeletal and heart muscle, that encodes a truncated protein of only the amino-terminal 181 residues (Futatsugi et al., 1998). It shares the splice acceptor site with the SIm2– variant.

Structure of IP3 receptor

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Domain Structure

IP3R forms homo- or hetero-tetrameric Ca2+ channels with six transmembrane domains. Among members of the ion channel family, IP3R is unique in its localization on intracellular Ca2+ stores and in its dual regulation of channel opening by two second-messengers, IP3 and Ca2+. Three IP3R isoforms and the splice variants show differences in ligand affinity, their regulatory factors, and expression or localization patterns, though their functional domain structures are apparently nearly the same. Herein, the domain structure of the best-characterized mouse IP3R1 (mIP3R1) is described as a representative (> Figure 24-1).

3.1 Five-Domain Structure Model mIP3R1 (SI+, SII+, SIII) is a 2749 amino acid polypeptide (calculated Mr = 313 kDa) and is structurally divided into three parts: a large amino (N)-terminal cytoplasmic region (83% of the receptor molecule), a six membrane-spanning domain, and a short C-terminal cytoplasmic tail (Michikawa et al., 1994). The bulky N-terminal cytoplasmic region is further divided by the ligand binding region into three domains. The five functional domains are called as the N-terminal coupling domain, ligand binding domain, internal coupling domain, transmembrane domain, and gatekeeper domain (> Figure 24-1) (Uchida et al., 2003).

3.1.1 N-Terminal Coupling Domain The N-terminal coupling domain (residues 1–225 of mIP3R1) is also called the ‘‘Suppressor domain’’ because it functions as the suppressor for IP3-binding to the ligand binding domain. Deletion of these residues results in significant enhancement of IP3 binding (more than tenfold higher affinity for IP3 than wild-type mIP3R1) (Uchida et al., 2003). Interestingly, the deletion mutant channel has no Ca2+ release activity despite having high IP3 affinity and is likely to retain the normal folding structure investigated by limited trypsin-digestion analysis (Uchida et al., 2003). This domain is therefore the key regulator of channel activity, via coupling with both the ligand binding domain and the channel-forming transmembrane domain. The crystal structure of the N-terminal coupling domain was unveiled at 1.8 A˚ resolution (> Figure 24-2a, b) (Bosanac et al., 2005). The structure contains a head domain forming the b-trefoil fold and an arm domain possessing a helix-turn-helix structure that protrudes from the globular head domain. Site-directed mutagenesis studies on the N-terminal coupling domain of mIP3R1 showed the residues (indicated in red in > Figure 24-2f ) on the conserved surface of the head subdomain to be critical for the suppression of IP3 binding (> Figure 24-2e) (Bosanac et al., 2005). Without the N-terminal coupling domain, the ligand-binding domain of all three IP3R isoforms shows the same high affinity for IP3 (see > Section 5) (Iwai et al., 2007). The isoform-specificity of IP3 affinity is therefore determined by combination with the N-terminal coupling domain (> Figure 24-3a). The N-terminal coupling domain is a binding target of numerous proteins (> Figures 24-1a and > 24-2f ), including CaM (Adkins et al., 2000; Sienaert et al., 2002), CaBP1 (Yang et al., 2002; Kasri et al., 2004), Homer (Tu et al., 1998), RACK1 (Patterson et al., 2004b), and partially Na+/K+-ATPase (Zhang et al., 2006). These proteins regulate the Ca2+ release activity of IP3R or regulate the interaction with cytoskeletal proteins to possibly communicate with plasma membrane proteins.

3.1.2 Ligand Binding Domain The ligand binding domain (residues 226–578 of mIP3R1) is also called the ‘‘IP3 binding domain’’ or ‘‘IP3binding core domain’’. Deletion mutagenesis indicated that IP3R binds IP3 within the N-terminal 650 amino acids independently of the tetramer formation (Mignery and Su¨dhof, 1990; Miyawaki et al., 1991),

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and the region was further narrowed to residues 226–578 of mIP3R1 (Yoshikawa et al., 1996). This region is a necessary and sufficient minimum region for specific IP3 binding, forming an ‘‘IP3-binding core’’. IP3 has three phosphate moieties in the equatorial positions 1, 4, and 5 of the inositol ring. Binding competition and Ca2+ release experiments using various synthetic analogues showed the positional distribution of the phosphate groups to be critical for ligand activity (Berridge and Irvine, 1984; Nahorski and Potter, 1989; Hirata et al., 1990; Wilcox et al., 1998). It was envisaged that the IP3-binding site has a pocket of positive charges facilitating ionic interaction with phosphate groups. The residues Arg265, Lys508, and Arg511 on the IP3-binding core region were found to be critical positively-charged residues for IP3 binding (Yoshikawa et al., 1996). The domain structure in complex with IP3 was uncovered at a 2.2 A˚ resolution (> Figure 24-2c–d) (Bosanac et al., 2002). The asymmetric, boomerang-like structure consists of an N-terminal b-trefoil domain, a hinge region (residues 435–437 of mIP3R1), and a C-terminal a-helical domain containing an armadillo repeat-like fold. The cleft formed by the b and a domains exposes a cluster of basic residues (> Figure 24-2d), and 11 amino acid residues, including mutationally determined Arg265, Lys508, and Arg511, on the cleft coordinate the three phosphoryl groups of IP3 (Bosanac et al., 2002). Crystal structure analysis also clarified that the head subdomain of the N-terminal coupling domain and the b subdomain of the ligand binding domain are both folded in the b-trefoil structure. The fold consists of six two-stranded b-hairpins, three of which make up a barrel while the other three are in a triangular array that caps the barrel. The two b-trefoil structures superimpose well despite the low amino acid sequence identity between them (Bosanac et al., 2005). Either Ca2+ or CaM binding to the N-terminal 600 amino acid residues reduces the binding of IP3 (Sienaert et al., 2002). Two CaM binding sites in the N-terminal coupling domain, residues 49–81 and 106– 128 of mIP3R1, and two Ca2+ binding sites in the ligand binding domain, residues 304–381 and 378–450 of mIP3R1, were estimated using a series of partial recombinant peptides of mIP3R1 (> Figure 24-1a) (Sienaert et al., 1997, 2002). Although the Ca2+ binding sites were mapped on the ligand binding domain, this inhibition requires the N-terminal coupling domain. It suggests that Ca2+-binding on the ligand binding domain changes the interacting affinity to the N-terminal coupling domain and regulates the extent of ligand suppression. IP3 binding is also regulated by IRBIT, which binds to the ligand binding domain in a phosphorylation dependent manner (Ando et al., 2003, 2006). Ten of the 12 key amino acids for IP3-recognition in the ligand . Figure 24-3 Isoform-specific ligand-binding affinity of IP3R and critical residues for type3-specific IP3 affinity. (a) The apparent IP3 dissociation constants of the ligand-binding domain (Core) of the three IP3R isoforms with or without the N-terminal coupling domain (Sup) and the intrinsic dissociation constants of full-length IP3Rs (Full length) estimated in b. Among 11 loop segments (L1–L11) that connect b-strands of the N-terminal coupling domain structure, L3/L5/L7 or L5/L7/L8 is critical for type-3 IP3R specific IP3-binding affinity. A chimeric protein composed of type 1 Sup and type 3 Core shows higher IP3-binding affinity (Kd~66 nM) than the native type 3 Sup+ Core (Kd~163 nM). Replacement of three loops, L3/L5/L7 or L5/L7/L8, of the type 1 Sup with the IP3R3sequences exhibited the lower IP3-affinity, indistinguishable from that of the native type 3 Sup+ Core. On the ribbon diagram representation of the N-terminal coupling domain of mouse IP3R1, the critical loops, L3, L5, L7, and L8 for type-3-specific suppression, are depicted in green, yellow, orange, and magenta, respectively. The conserved residues essential for IP3-binding suppression (Figure > 24-2f) are depicted in red. On the surface representation, the type-3-specific amino acid residues on L3, L5, L7, and L8 are depicted in green, yellow, orange, and magenta, respectively. (b) Different IP3-binding affinities and cooperativities of three types of tetrameric IP3R channels. Values are determined from the curve fitting analysis of the equilibrium IP3-binding activity graph (left) (Iwai et al., 2005). A tetrameric IP3R-channel has four IP3-binding sites. The K value is the intrinsic association constant of the vacant IP3-binding site in a tetrameric receptor in which no IP3-binding site is occupied. IP3 binding to a homotetrameric IP3R1 channel occurs in a non-cooperative manner. On the other hand, IP3R2 channels exhibit both negative and positive cooperativities and IP3R3 channels exhibit a negative cooperativity in all IP3-binding steps (see the online version at www.SpringerLink.com)

. Figure 24-3 (continued)

Structure of IP3 receptor

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binding domain participate in binding to IRBIT, and IRBIT suppresses the activation of IP3R by competing with IP3 as an endogenous ‘‘pseudoligand’’ (Ando et al., 2006). IRBIT is also presumed to be a ‘‘third’’ messenger, released from IP3R in response to the production of IP3, and to regulate cell functions through downstream target molecules (Shirakabe et al., 2006).

3.1.3 Internal Coupling Domain The internal coupling domain (residues 579–2275 of mIP3R1) is also called the ‘‘Modulatory domain’’ or ‘‘Transducing domain’’ since various binding factors (> Figure 24-1a) to this region modulate IP3R channel activity, and this domain is involved in the signal transduction of IP3-binding to channel gating (Mignery and Su¨dhof, 1990; Uchida et al., 2003). This domain shares the least homology among the three IP3R isoforms, suggesting a regulatory variation in different subtypes. Little information has been obtained about the structure of the internal coupling domain. Previous amino acid sequence analyses suggested a high helical propensity and the possibility of an armadillo-like fold continuing from a part of the ligand binding domain, residues 438–1740 of mIP3R1 (Ponting, 2000; Bosanac et al., 2002). From the threedimensional structure of IP3R (> Figure 24-4a), the internal coupling domain is supposedly folded into a compact but porous globular structure and various factors access the region to regulate IP3R activity (Sato et al., 2004). One glutamate residue, corresponding to Glu2100 of mIP3R1 in the internal coupling domain, is regarded as a component of a Ca2+sensor that regulates the Ca2+ sensitivity of IP3R (Miyakawa et al., 2001). This residue was found by sequence alignment between IP3R and the ryanodine receptor (RyR), in which the corresponding residue is important for the Ca2+ sensitivity of RyR channel activity (Chen et al., 1998;

. Figure 24-4 3D structures of mIP3R1. (a and b) Highest resolution EM structure of mIP3R1, determined in the absence of IP3 and Ca2+, and viewed from the cytosol, side, and ER lumen, respectively in a (adapted from Sato et al., 2004). ˚ . The crystal structure of The putative transmembrane zone is indicated by cyan square brackets. Bar, 100 A the ligand binding domain, represented both by a ribbon diagram and a space-filled model, is modified and fitted into the density map of the whole IP3R channel structure in b (adapted from Sato et al., 2004). (c) Global structural change from square-state (left) to windmill-state (right) of mIP3R1 resulting from the presence of Ca2+. The crystal structure of the ligand binding domain (blue) in complex with IP3 (red) is modeled in both conformational states (adapted from Hamada et al., 2003) (see the online version at www.SpringerLink.com)

Structure of IP3 receptor

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Du and MacLennan, 1998). Substitution of Glu2100 to Asp results in a tenfold decrease in Ca2+ sensitivity without affecting either IP3 sensitivity or the maximal rate of Ca2+ release activity (Miyakawa et al., 2001), and the mutation also shifts the peak of the IP3R bell-shaped Ca2+ dependence to a higher Ca2+ concentration (Tu et al., 2003). The Glu2100 site is not included among the previously determined putative Ca2+ binding sites (> Figure 24-1a) (Sienaert et al., 1996, 1997). The Glu2100 mutation might allosterically change the Ca2+ affinity of Ca2+-binding sites.

3.1.4 Transmembrane Domain The transmembrane domain (residues 2276–2589 of mIP3R1) (> Figure 24-1) has six transmembrane helices, M1 to M6 (Yoshikawa et al., 1992; Michikawa et al., 1994), which share similar topology with other cation channels, including the super family of voltage-gated ion channels, cyclic nucleotide-gated ion channels, and trp family of ion channels. This domain is responsible for the tetramer formation of IP3R (Mignery and Su¨dhof, 1990), and the region from M5 to M6 is the key element stabilizing the IP3R tetramer (Joseph et al., 1997; Galvan et al., 1999; Galvan and Mignery, 2002). The linker segment between M5 and M6 (residues 2463–2569 of mIP3R1) has a relatively long luminal loop and was suggested to be the poreforming region of IP3R based on the structural analogy with voltage- and second messenger-gated ion channels (Michikawa et al., 1994). Pore-formation with this loop was also proposed since deletion of the M1–M4 region produced a constitutively open channel (Ramos-Franco et al., 1999). The M5–M6 loop can be divided into two regions, V (variable) and C (conserved) (> Figure 24-1b) (Michikawa et al., 1994). The first half (V) has highly divergent sequences according to the IP3R isoform, suggesting that this region is involved in isoform-specific regulation. On the other hand, the second half (C) is almost completely conserved among isoforms and species of the IP3R family. In the C region, a poreforming helix and a selectivity filter were estimated to exist based on the sequence and secondary structure similarity with RyR and a potassium channel, KcsA (Michikawa et al., 1994; Doyle et al., 1998; Shah and Sowdhamini, 2001; Williams et al., 2001). Mutations of Asp2550 in the estimated selectivity filter (residues 2546-GGVGD-2550 of mIP3R1), to Ala or Asn, resulted in channel impermeability to Ca2+, and substitution to Glu yielded channels lacking selectivity for Ca2+ over K+ (Boehning and Joseph, 2000a; Boehning et al., 2001). On the other hand, Val2548 is critical for ion conductance control, since its mutation to Ile leads to channels with higher K+ permeation but the same Ca2+ selectivity as wild-type channels (Boehning et al., 2001). The IP3R channel activity is regulated by not only cytosolic factors but also ER luminal proteins. ERp44, a member of the thioredoxin family, binds to the V region of the M5–M6 loop in a type 1 IP3R specific manner (Higo et al., 2005). The interaction requires cysteine residues in the loop and is dependent on pH, Ca2+ concentration, and redox state. ERp44 negatively regulates the Ca2+ release activity of IP3R. On the other hand, chromogranin A and B, members of the Ca2+ storage protein family, which have low affinity and high binding capacity for Ca2+, bind to the C region of the M5–M6 loop of all three IP3R isoforms (Yoo and Lewis, 1998, 2000; Kang et al., 2007) and activate the Ca2+ release activity of IP3R (Thrower et al., 2003). Chromogranin A binds to IP3R restrictedly around pH 5.5, while chromogranin B binds over a wider pH range (Yoo et al., 2000). Another loop of the transmembrane domain, the M4–M5 linker, is directed to the cytosol and is essential for channel gating (see > Section 3.3).

3.1.5 Gatekeeper Domain The gatekeeper domain (residues 2590–2749 of mIP3R1) is also called the ‘‘C-terminal coupling domain’’. The secondary structure of this last 160 residues is little known, but is suggested to be a highly helical structure (Bosanac et al., 2004; Schug and Joseph, 2006). An important role of this domain in channel gating was theorized from the observation that binding of monoclonal antibody to the residues 2736–2747

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of mIP3R1 (Nakade et al., 1991) and the expression of the C-terminal GFP-fused IP3R (Nakayama et al., 2004) suppress channel activity. Furthermore, mutation of a highly conserved residue, Cys2613 of mIP3R1, resulted in a non-functional Ca2+ channel (Uchida et al., 2003). Several regulatory proteins have been reported to bind to the gatekeeper domain (> Figure 24-1a) (Patterson et al., 2004a; Foskett et al., 2007).

3.2 Trypsin Digestion Pattern Model The mIP3R1 protein complex is digested into five major fragments (I–V) by limited (controlled) trypsinization of microsomal fractions from mouse cerebellum (> Figure 24-1a) (Yoshikawa et al., 1999). Fourfragments, I–IV, are derived from the N-terminal cytoplasmic region and the C-terminal fragment V includes the membrane-spanning channel region. The boundary of fragments I and II resides on the loop between the b6- and b7-strands of the ligand-binding domain (> Figure 24-2c). The intrinsic flexibility of this region is presumed not only from the high accessibility of trypsin, but also the invisible structure of the loop in crystal analysis. All five fragments are well folded, not easily trypsinized, and form stable noncovalent interactions with each other, such that they coimmunoprecipitate with the C terminus-specific monoclonal antibody after trypsinization and solubilization with 1% Triton X-100. Surprisingly, the five major fragmented mIP3R1 channels retain significant IP3-induced Ca2+ release activity (Yoshikawa et al., 1999).

3.3 Head-To-Tail Interaction Model How does the binding of IP3 at the N terminus of IP3R gate the opening of the channel pore some 2,000 amino acids away at the C terminus? The necessity of both the N-terminal coupling domain and the C-terminal gatekeeper domain has been suggested (see > Section 3.1). Limited trypsinization studies proposed an interaction between the N- and C-termini of IP3R, though the other fragments interacting noncovalently might intervene (see > Section 3.2) (Joseph et al., 1995; Yoshikawa et al., 1999). Crosslinking and pull-down assays defined the direct interaction between residues 1–340 in the N-terminus and residues 2418–2749, including the downstream region of the transmembrane segment M4 to the C-terminus, with association occurring in an intersubunit manner (Boehning and Joseph, 2000b). Truncation mutagenesis further clarified the loop between M4–M5 linker (> Figure 24-1b), to a crucial region for the head-to-tail interaction and to be required for channel gating (Schug and Joseph, 2006). A region of the C-terminal tail (residues 2694–2721), predicted to be a coiled-coil, is also required for channel activity and stabilization of the head-to-tail interaction (Schug and Joseph, 2006). Head-to-tail interaction has been observed in cyclic nucleotide-gated ion channels (Varnum and Zagotta, 1997; Gordon et al., 1997; Rosenbaum and Gordon, 2002) and voltage-gated and inwardly rectifying potassium channels (Schulteis et al., 1996; Tucker and Ashcroft, 1999). The importance of the M4–M5 linker (also called the S4–S5 linker) has been emphasized in the gating mechanism of voltage-gated channels (Long et al., 2005a, b). The voltage-induced movement of the charged M4 helix has been proposed to displace the M4–M5 linker and thereby permit the M6 helix bundle to separate, thereby allowing ion conduction. Similarly, in inwardly rectifying potassium channels, KirBac, a homologous M4–M5 linker structure referred to as a ‘‘sliding helix’’ has been suggested to play a critical role in channel gating (Kuo et al., 2005). IP3R has no homologous sequences with the M4 helix of voltage-gated channels, and, in fact, the cerebellar IP3R reconstituted into the planar lipid bilayer was only weakly voltage dependent (Watras et al., 1991), while the conformational change directed by IP3-binding might drive the movement of the M4–M5 linker through the head-to-tail interaction (> Figure 24-1b). Consequently, the coupling among the N-terminal coupling domain, the M4–M5 linker, and the C-terminal gatekeeper domain of IP3R is quite important for propagation of the IP3-binding signal to channel gating, and the internal coupling domain also supports the signal transfer via an unidentified mechanism (Uchida et al., 2003).

Structure of IP3 receptor

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Three-Dimensional Structure

Clarification of the three-dimensional (3D)-structure of IP3R has been sought for understanding channel gating mechanism, though structural studies are enormously difficult because the IP3R channel is a gigantic membrane protein, which has a molecular mass of 1,200 kDa for the tetrameric complex. Earlier negative stain electron microscopy (EM) studies of solubilized and purified receptors from microsomal fractions (Chadwick et al., 1990; Maeda et al., 1990) revealed a square or pinwheel-like 2D-structure with fourfold symmetry. Using quick-freeze deep-etch replica EM, the array images of native IP3R on the endoplasmic reticulum membranes of cerebellar Purkinje neurons (Katayama et al., 1996) were clarified. In 2002 and 2003, several groups reported the 3D-structure of IP3R using single particle reconstruction from cryo-EM images of isolated receptors (Jiang et al., 2002a; da Fonseca et al., 2003; Hamada et al., 2003; Serysheva et al., 2003), though there are many discrepancies in size (e.g., the lateral dimension varies from 150 to 250 A˚) and in shape, possibly because of the differences in methods of sample preparation and data analysis. Using an originally developed cryo-EM equipped with a helium-cooled specimen stage and an automatic particle picking system, the density map of IP3R was obtained at high resolution (15 A˚) through image analysis of about 10,000 particles (> Figure 24-4a) (Sato et al., 2004). The shape of the mIP3R1 with an overall height of 231 A˚ is reminiscent of a hot-air balloon, with the spherical cytoplasmic domain (diameter of 175 A˚) and the square-shaped luminal domain (side length of 96 A˚). The structure has multiple internal cavities and contains four prominent L-shaped densities on the top part of the cytoplasmic domain. The L-shaped densities are estimated to be the ligand-binding domain because only this region could accommodate the X-ray structure of the ligand-binding domain with some modification (> Figure 24-4b) (Sato et al., 2004). The assignment of the ligand-binding domain on the 3D structure is consistent with mapping using heparin-gold labeling (Hamada et al., 2002) and the density fit (Serysheva et al., 2003), but disagrees with the assignment to the nearest region of the fourfold axis (da Fonseca et al., 2003), which was inferred from an earlier ligand-binding study using synthetic IP3 dimers suggested by the ligand binding domain within a tetrameric receptor which is not more than 20 A˚ apart (Riley et al., 2002). The aforementioned 3D-structures were all obtained from purified IP3R prepared in a Ca2+-free buffer. Remarkable 3D-structural change of IP3R was observed in Ca2+-containing buffer under negative stain EM (> Figure 24-4c) (Hamada et al., 2002, 2003). The structure of mIP3R1 without Ca2+ has a ‘‘mushroomlike’’ appearance consisting of a large square-shaped head and a small channel domain. By the addition of Ca2+, less than 100 nM free Ca2+, the structure changes to the ‘‘windmill-like’’ form. This global change might be partially due to the uranyl acetate staining, since the very low pH of the solution may cause artificial structural changes in the flexible molecules, though this result first clarified the structural change with Ca2+. In contrast, IP3 scarcely affected the conformational states (Hamada et al., 2002). Recently, the IP3R structure was visualized using atomic force microscopy and the first 3D representation of individual IP3R particles in aqueous solution was provided (Suhara et al., 2006).

5

Functional Properties of IP3R Isoforms

Mammalian IP3R isoforms, IP3R1, IP3R2, and IP3R3, share 60–70% identity (see > Section 2.1) and each isoform exhibits a unique expression pattern. IP3R1 is predominant in the central nervous system, especially in cerebellar Purkinje cells (Maeda et al., 1988; Worley et al., 1989; Furuichi et al., 1993), while most tissues express at least two and often all three IP3R isoforms at different ratios (Taylor et al., 1999). Heterotetramer formation among distinct IP3R isoforms (Monkawa et al., 1995; Onoue et al., 2000) augments IP3R channel functional diversity in cells. To understand the diverse regulatory roles of IP3R in Ca2+ signaling, it is important to characterize both the conserved and isoform-specific properties of IP3R. IP3 binding affinities of three IP3R isoforms were compared under identical experimental conditions (Iwai et al., 2005). Recombinant mouse IP3Rs expressed in Spodoptera frugiperda (Sf9) cells show IP3 binding affinity with the order of IP3R2 (the dissociation constant Kd = 5.9 nM) > IP3R1 (28.6 nM) > IP3R3

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Structure of IP3 receptor

(294.0 nM) under Ca2+-chelated conditions (> Figure 24-3a, Full length). This relative order is consistent with the result of the N-terminal domain (600 residues) of rat and mouse IP3Rs (Newton et al., 1994; Iwai et al., 2007; > Figure 24-3a, Sup + Core). On the other hand, without the N-terminal coupling domain (see > Section 3.1), the ligand binding domain of all three isoforms exhibits a nearly identical IP3-binding affinity (Kd2 nM) (> Figure 24-3a, Core) (Iwai et al., 2007). The N-terminal coupling domain can physically interact with the IP3-binding core domain (Bultynck et al., 2004). These results suggest that the interaction between the N-terminal coupling and IP3-binding domains produces the isoform-specificity for IP3 binding. In IP3R3, the amino acid residues that contribute to generate isoform-specific IP3-affinity were identified within the N-terminal coupling domain (> Figure 24-3a) (Iwai et al., 2007). Tetrameric channel formation further increases the diversity of IP3 binding properties because of ligand binding cooperativity. IP3 binding to a homotetrameric IP3R1 channel occurs in a non-cooperative manner, though an IP3R2 (SIm2+) channel exhibits both negative and positive cooperativities and an IP3R3 channel exhibits negative cooperativity (> Figure 24-3b) (Iwai et al., 2005). IP3R isoforms have well-conserved ion permeability properties among different species. The IP3R channel is a divalent-selective cation channel exhibiting a selectivity Ca2+: K+ ratio of 6, with relatively little selectivity among different divalent cations (Foskett et al., 2007). Under identical experimental conditions, Tu et al. (2005a, b) compared the single-channel behavior of recombinant rat IP3Rs, which were expressed in Sf9 cells and reconstituted into planar lipid bilayers. Using 50 mM Ba2+ as the current carrier, all three IP3Rs show 1.9 pA unitary current and 80 pS single-channel conductance. Under optimal conditions, the three IP3Rs show comparable single-channel properties of the open probability, the mean open dwell time, and the mean closed dwell time. Drosophila melanogaster IP3R displayed single-channel properties similar to those of these mammalian IP3R isoforms (Srikanth et al., 2004), indicating that these major functional properties of IP3R are conserved during evolution. Isoform specificity of the single-channel open probability is generated by numerous factors, including ATP and coagonists, IP3 and Ca2+. IP3 sensitivity was determined to be in the order of IP3R2 > IP3R1 > IP3R3 under conditions of pCa 6.7 (cytoplasmic side) and 0.5 mM ATP (Tu et al., 2005b). The relative order is consistent with the IP3-binding affinity mentioned above and with the functional analysis of IP3R isoforms (Miyakawa et al., 1999). ATP also potentiates IP3R channel activity, although ATP is not necessary for channel gating (Taylor and Putney, 1985; Meyer et al., 1988; Bezprozvanny and Ehrlich, 1993) and ATP alone is not sufficient to open the IP3R channel (Ferris et al., 1990; Bezprozvanny et al., 1991; Iino 1991). Tu et al. (2005a) reported that the ATP-sensitivity is in the order of IP3R1 > IP3R3 and that IP3R2 is ATP independent under conditions of pCa 6.7 (cytoplasmic side) and 2 mM IP3, in agreement with previous functional studies (Miyakawa et al., 1999; Maes et al., 2000). High ATP-sensitivity of IP3R1 most probably results from the unique high-affinity ATP binding site (ATPA site in > Figure 24-1a) (Maes et al., 2001; Tu et al., 2002; Tu et al., 2005b). Modulation of IP3R open probability by cytosolic Ca2+ is one of the most fundamental IP3R properties. At low Ca2+ concentrations, Ca2+ acts as coactivator of the IP3R, while at higher Ca2+ concentrations IP3R is inhibited by Ca2+ (Iino, 1990; Bezprozvanny et al., 1991; Finch et al., 1991). This property of IP3R, called bell-shaped Ca2+ dependence, is essential for the generation of Ca2+ waves and Ca2+ oscillations (De Young and Keizer, 1992; Lechleiter and Clapham, 1992; Berridge, 1993; Parker et al., 1996). Despite earlier controversy (Hagar et al., 1998; Ramos-Franco et al., 1998, 2000), there is now agreement that all IP3R isoforms display bell-shaped Ca2+ dependence in a physiological range of Ca2+ concentrations (Taylor and Laude, 2002; Tu et al., 2005a; Foskett et al., 2007). Eight putative Ca2+-binding sites of IP3R1 have been determined using a series of partial recombinant peptides (> Figure 24-1a), to date, though which sites are responsible for the stimulatory or inhibitory Ca2+-binding remains unknown. IP3 fine tunes the bell-shaped Ca2+ sensitivity of IP3R (Kaftan et al., 1997; Mak et al., 1998, 2001; Foskett et al., 2007). At a low IP3 concentration, IP3R channels are quite sensitive to Ca2+ inhibition and channel activity is inhibited before achieving full activation, while a higher IP3 concentration relieves the Ca2+ inhibition and would promote greater diffusion of local Ca2+ signals. Thus, IP3R single-channel properties are regulated by its ligands in a complicated manner, and the isoform-specific sensitivity for the ligands provides diversity in cytoplasmic Ca2+ signals as reported for isoform-specific Ca2+-signaling patterns in cells (Miyakawa et al., 1999; Hattori et al., 2004).

Structure of IP3 receptor

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Concluding Remarks

Recent progress in structural and functional studies of IP3R provides an abundance of information about molecular properties as a Ca2+ channel. Physiological diversity of IP3-induced Ca2+ signals is generated by combinations of several IP3R isoforms and also by various interacting factors. For fully understanding of IP3R functions, future studies should clarify how IP3-induced conformational changes propagate through IP3R to open the channel pore and how the modulators regulate the processes. It is also important to elucidate the functional relationships among various channels and pumps, which reside on both intracellular and plasma membranes, regulating cytoplasmic Ca2+ concentrations.

Acknowledgments We gratefully acknowledge Drs. Mitsuhiko Ikura, Chikara Sato, Yoshinori Fujiyoshi, and their laboratory members for their collaboration in the structural analyses. We also thank the many colleagues whose names are cited in the references. The research in our laboratory is supported by grants from the Japan Science and Technology Agency (KM) and by Grants-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology of Japan (KM).

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Insights into the Three-Dimensional Organization of Ryanodine Receptor

L. G. D’Cruz . C. C. Yin . A. J. Williams . F. Anthony Lai

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 464

2 2.1 2.2 2.3 2.4 2.5 2.6 2.7

Ryanodine Receptors in Human Pathology and Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 464 Catecholaminergic Polymorphic Ventricular Tachycardia (CPVT) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 464 Arrhythmogenic Right Ventricular Cardiomyopathy (ARVC2) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 465 Malignant Hyperthermia (MH) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 465 Central Core Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 465 Myasthenia Gravis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 466 Alzheimer’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 466 Structural Information to Understand Disease Pathophysiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 466

3 cryo-EM Topology of the RyR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 467 3.1 Cytoplasmic Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 467 3.2 Transmembrane Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 469 4 4.1 4.2 4.3 4.4 4.5 4.6 4.7 4.8

RyR Domain Organization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 469 MIR Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 469 SPRY Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 470 EF Hand Domains and Ca2+ Binding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 471 Transmembrane Domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 473 FKBP Binding Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 474 RyR Phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 476 DHPR-RyR Interaction Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 476 RyR Binding to Accessory Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 477

5

Comparison of RyR Isoforms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 478

6

Open and Closed States of RyR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 478

7

Interaction Between RyRs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 479

8

Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 479

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2009 Springer ScienceþBusiness Media, LLC.

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Insights into the three-dimensional organization of ryanodine receptor

Abstract: The ryanodine receptor encompasses a high-conductance calcium channel that plays a critical role in intracellular calcium signalling and its dysfunction has been implicated in numerous cardiovascular and neuromuscular disorders. Here, we discuss the various pathologies associated with aberrant ryanodine receptor function and our current understanding of the intricate topological organisation of this the largest known ion channel complex. List of Abbreviations: APP, amyloid precursor protein; ARVC, arrhythmogenic right ventricular cardiomyopathy; CaMKII, Ca2+/calmodulin-dependent kinase; CICR, Ca2+-induced Ca2+ release; CaM, calmodulin; CPVT, catecholaminergic polymorphic ventricular tachycardia; CCD, central core disease; CPK, conventional protein kinase; cryo-EM, cryo-electron microscopy; CRISP, cysteine-rich secretory protein; Kd, dissociation constant; DR1, DR2, and DR3, divergent regions 1, 2, and 3; FAD, familial Alzheimer’s disease; FKBP, FK506 binding protein; IpTxa, imperatoxin A; IP3R, inositol trisphosphate receptor; ITC, isothermal titration calorimetry; MH, malignant hyperthermia; MIR, mannosyltransferase, IP3R and RyR; PS1, presenilin 1; PKA, protein kinase A; RyR, ryanodine receptor; RIH, RyR and IP3-homology; SPRY, splA and ryanodine receptor; SOCS, suppressor of cytokine signaling; TM, transmembrane

1

Introduction

The ryanodine receptors (RyRs) belong to a family of intracellular calcium release channels that include the inositol trisphosphate receptors (IP3Rs) that are also reviewed elsewhere in this book. Historically, it is this channel’s ability to specifically bind to the neutral plant alkaloid, ryanodine, to which the receptor owes its name. Ryanodine binds to the high (nanomolar) affinity site of the RyR molecule, locking the receptor in an open state whose conductance is 40% that of the open state with Ca2+ as the permeant ion. RyRs are the largest known integral membrane ion channel proteins, comprising subunits of 5,000 amino acids that form homotetrameric structures with an approximate molecular weight of 2,300 kDa. There are three mammalian isoforms; RyR1 is predominantly found in skeletal muscle, RyR2 is found mainly in muscles of the heart whereas RyR3 was initially identified in brain tissues but has since been found expressed at low levels in a variety of other tissues. Mutations in the gene encoding the RyR molecule are translated into an aberrant protein with dysfunctional channel properties that contribute to a number of disease states. Understanding the structural properties of this enormous protein would lead to a better understanding of the way in which the RyR functions as a calcium channel. It follows then, that understanding the biology of cell calcium regulation via the RyR would help in developing therapeutic approaches to treat the disease states that are attributed to this molecule.

2

Ryanodine Receptors in Human Pathology and Disease

2.1 Catecholaminergic Polymorphic Ventricular Tachycardia (CPVT) One of the most important features of the RyR in medicine and health is that channel dysfunction is linked to an early-onset cardiac disease in which affected individuals are usually diagnosed during childhood or adolescence and 30–50% of the affected patients succumb to sudden death by the age of 30 (Fisher et al., 1999; Swan et al., 1999). This RyR2-mediated cardiac channelopathy that can lead to sudden death is an arrhythmogenic disorder known as catecholaminergic polymorphic ventricular tachycardia (CPVT), a condition characterized by stress-induced bi-directional or polymorphic ventricular tachycardia (VT). This form of polymorphic VT frequently degenerates into ventricular fibrillation, which culminates in sudden cardiac death in the absence of structural heart disease. CPVT is a genetic disease and has an autosomal-dominant mode of inheritance [OMIM 604772]. Linkage studies and direct sequencing have identified mutations in the human cardiac ryanodine receptor gene (hRyR2) on chromosome 1q42-q43 of individuals with CPVT (Swan et al., 1999; Laitinen et al., 2001; Priori et al., 2001).

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To date, over 80 distinct mutations identified in RyR2 have been associated with at least two forms of cardiac arrhythmia, CPVT and arrhythmogenic right ventricular dysplasia type 2 (ARVD2). Most of these mutations are located in the N-terminal region (between amino acids 164–466), central region (2,246–2,504), and the C-terminal region (3,778–4,959) of RyR2 (Thomas et al., 2007).

2.2 Arrhythmogenic Right Ventricular Cardiomyopathy (ARVC2) Arrhythmogenic right ventricular cardiomyopathy (ARVC) is a progressive debilitating heart disease characterized by fibrofatty replacement of the myocardium, primarily affecting the right ventricle but may also involve the left ventricle with disease progression (Horimoto et al., 2000). Those affected by this disorder include the young and athletes, who are predisposed to ventricular tachycardia (VT) and sudden death (Marcus et al., 1982; Thiene et al., 1988). There are subtle variations within the disease associated with three different genes that specify its autosomal dominant mode of inheritance. ARVD2 caused by mutations in the RyR2 gene (Tiso et al., 2001), ARVD8 caused by mutations in the gene for desmoplakin (Rampazzo et al., 2002) and ARVD9 caused by mutations in plakophilin-2 (Gerull et al., 2004). Six other variants of the disease (ARVD1, ARVD3, ARVD4, ARVD5, ARVD6 and ARVD7) have been characterized and their chromosomal loci mapped, however the causative genes have not yet been identified. It is important to know the causative genes for diseases, especially in ARVC/D since the clinical diagnosis of ARVD is complicated by the diversity of phenotypic features and the weakness of clinical tools in identifying gene carriers during the early ‘‘concealed’’ phase of the disease.

2.3 Malignant Hyperthermia (MH) Another RyR-linked pathological condition is an acute, life-threatening disease known as malignant hyperthermia (MH) or ‘‘malignant hyperpyrexia due to anesthesia.’’ These patients develop very high fever, muscle rigidity, breakdown of muscle fibers (rhabdomyolysis), increased acid levels in the blood and other tissues (acidosis) and a rapid heart rate when exposed to certain drugs used in general anesthesia. The most common trigger that has been implicated in cases of MH are the class of drugs known as ‘‘volatile anesthetics,’’ usually preferred by surgical anesthetists as they are generally safe for patients of all ages including pregnant women. They are generally odorless, reasonably pleasant to inhale, not metabolized and rapid in onset and offset of anesthesia. In individuals with MH, these inhalational anesthetics trigger a sudden increase in the oxidative metabolism of skeletal muscle to a point where the body is unable to adequately deliver oxygen and remove carbon dioxide, leading to catastrophic failure of body temperature regulation (hence hyperthermia) and this results in circulatory collapse and death. Almost 50–70% of cases of MH are caused by mutations in the skeletal muscle isoform of the ryanodine receptor (RyR1) (Gillard et al., 1991; Galli et al., 2006). In cases where there is a suspected familial predisposition to MH, patients who would be subjected to procedures involving general anesthesia would be given the diagnostic ‘‘caffeine-halothane-contracture test.’’ A small muscle biopsy from the patient, removed under local anesthesia, is bathed in solutions containing caffeine or halothane and MH-specific contraction is detected by microscopy.

2.4 Central Core Disease Mutations in RyR1 also result in a skeletal muscle disorder known as central core disease (CCD). This condition typically presents itself in infancy with marked hypotonia (low muscle tone) and a delay in normal motor development skills such as walking, sitting or standing, with pronounced proximal weakness in the hip girdle and upper legs. A frequent complication in CCD is susceptibility to MH, and the pathophysiological overlap in these conditions may be a consequence of sharing of the same allelic mutational sites in the RyR1 gene. The CCD muscle fibers appear to be microscopically disorganized at the centre, forming distinctive areas termed ‘‘cores’’ (hence the name of this disease). These CCD biopsy

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observations may resemble the myocyte disarray that is observed in hypertrophic cardiomyopathy (Davison et al., 2000), however patients with this form of cardiomyopathy do not present with profound skeletal muscle weaknesses or the other complications manifest in CCD (Jungbluth, 2007).

2.5 Myasthenia Gravis Myasthenia gravis (MG) is a debilitating neuromuscular disease characterized by fluctuating levels of muscle weakness, fatigue and thymomas. Individuals with MG have been shown to have high circulating auto-antibodies to acetylcholine receptors as well as to the RyR (Mygland et al., 1993). Although mutations in RyRs have not been linked to cases of MG, the circulating RyR auto-antibodies appear to interfere with the fundamental role of muscle RyR1 in the vital physiological function known as excitation-contraction coupling (Nakata et al., 2007).

2.6 Alzheimer’s Disease Neurodegeneration observed in Alzheimer’s disease has been linked to intracellular accumulation of misfolded proteins (amyloid plaques) and dysregulation of intracellular Ca2+ homeostasis. Aberrant intracellular Ca2+ regulation correlates with increased expression levels of RyR channels in neuronal cells that are thought to arise from transcriptional upregulation by the intracellular domain proteolytic fragment of amyloid precursor protein (APP). APP is a 4.2 kDa polypeptide of 28-amino acids that folds into a partial beta-pleated sheet structure and is implicated in cerebrovascular amyloidoses and in amyloid plaques associated with Alzheimer’s disease (OMIM: 104760). Proteolytic post-translational processing of APP by alpha-secretases generates a soluble amyloid protein, while beta- and gammasecretases generate APP components with amyloidogenic features. Familial Alzheimer’s disease (FAD) presenilin 1 (PS1) mutations result in enhanced Ca2+ responses to various stimuli, an increased predisposition of cells to undergo apoptosis, heightened capacitative Ca2+ entry and an increased gamma-secretase activity. APP proteolysis by gamma-secretase generates several fragments, including the APP-intracellular domain fragment (AICD), which has been shown to regulate IP3-mediated Ca2+ signaling via transcriptional mechanisms (Leissring et al., 1999a, b, 2001). In transgenic mouse models carrying presenilin mutations, attenuated Ca2+-induced Ca2+ release (CICR) through RyR has been observed (Stutzmann, 2007; Stutzmann et al., 2007).

2.7 Structural Information to Understand Disease Pathophysiology To understand the fundamental processes linking RyRs to disease pathways, it is vital to have a clear mechanistic insight of the Ca2+ signaling processes that RyRs mediate. From analysis of the RyR primary amino acid sequence we can glean some information as to where associated ligands and effectors bind. One can then hypothesise the effect of an RyR mutation on the binding of an effector, or how it alters channel function. However, proteins normally fold and adopt intricate 3-dimensional (3D) conformations and initial educated guesses from the primary amino acid sequence often require revision when the actual folding of the whole molecule is observed. X-ray crystallography is the method of choice for protein structure elucidation, however, protein crystals seldom demonstrate perfect periodicity and some of the diffraction is lost to diffuse scattering, thus complicating structure determination. Additionally, in order to obtain enough material for initial screens for crystallization conditions, one needs large amounts of the native protein. For the RyR, this would have to be obtained from muscle preparations isolated from a number of organisms, which is compounded by the issue of variations between protein preparations. Membrane proteins such as the RyRs are also notoriously difficult to crystallize because they exist partly in a highly hydrophobic membrane lipid environment. Thus, commonly used polar buffer solutions to

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solubilize protein preparations would cause aggregation of the proteins and may even effect partial unfolding. Due to these difficulties in protein crystallography of the RyR molecule, the structural information that we currently have is limited to cryo-electron microscopy (cryo-EM) data. The first 3D structures of RyR by cryo-EM and single-particle image processing were obtained from micrographs of frozenhydrated, detergent-solubilized RyR1s (Radermacher et al., 1994; Serysheva et al., 1995). In this article, RyR topology from cryo-EM analysis is described followed by putative RyR domain organization, based upon sequence comparison with related families of proteins in the SMART database (Schultz et al., 1998). Correlation of the RyR cryo-EM structural data with the primary amino acid sequence is considered together with recent advances in co-crystallization of RyR domains and accessory proteins. Homology modeling information on the transmembrane (TM) region is also discussed in combination with insights to its relevance in the overall function of the channel.

3

cryo-EM Topology of the RyR

The initial image reconstructions were obtained by two different approaches: the random conical reconstruction (Radermacher, 1994) and the angular reconstitution methods (Serysheva et al., 1995). In the random conical method, the specimen is tilted in the microscope to obtain many views of the RyR1 complex in different orientations, whose relative angles are defined by settings of the goniometer in the microscope. The angular reconstitution approach exploits random orientations of ice-embedded molecules imaged in the electron microscope without tilting the specimen holder. In this technique, the relative orientations of particles are computationally determined by searching their common line projections. The various methods in which cryo-EM data have been used to reconstruct 3D images have been reviewed elsewhere (Saibil, 2000). Although different methods of data processing have been used, the final results show similar topological surface maps of the RyR. If the analogy of a mushroom is used to describe the RyR topological surface, the majority of the protein i.e., the mushroom ‘‘head’’ corresponds to the cytoplasmic region (> Figure 25-1). When viewed from the cytoplasm, the RyR has the overall shape of a square prism with dimensions, 280  280  120 A˚. The ‘‘stalk’’ of the mushroom, represents the TM portion (> Figure 25-1). The assignment of the cytoplasmic and TM regions were deduced from examining electron micrographs of the RyR in situ, where they remain integrated in their natural membrane environment (Block et al., 1988; Saito et al., 1988). The clear fourfold symmetry is consistent with the RyR being a homotetramer (Lai et al., 1989). Recently, with the aid of advanced electron microscopes, the resolution of the 3D reconstructions have been improved to 10 A˚ and below using improved image processing algorithms (Ludtke et al., 2005; Samso et al., 2005; Serysheva et al., 2008).

3.1 Cytoplasmic Assembly The surface topology of the RyR cytoplasmic region appears to consist of ten or more discrete globular domains per subunit that are clearly resolvable due to their separation by solvent-accessible regions. Image reconstructions from both the Wagenknecht and Chiu groups show identical arrangement of the various globular domains although different methods of nomenclature are used by these two groups. The Wagenknecht group assign numerical labels to the domains whereas the Chiu group prefer descriptive names. Of the globular domains, the one labeled as domain ‘‘3’’ by the Wagenknecht group, or ‘‘handles’’ as described by the Chiu group, is the largest of the globular domains (> Figure 25-1). Domains 5–10 form the corners of the cytoplasmic assembly, these being described as ‘‘clamps’’ by the Chiu group (> Figure 25-1). A central 40–50 A˚ diameter solvent-filled pocket which connects to the TM region is surrounded by four ‘‘2’’ domains (see > Figure 25-1). The ‘‘clamps’’ or domains 5–10 appear to be connected to the receptor via three interactions: (1) the ‘‘handles’’ or domain ‘‘3’’ connect domains 5 and 9, (2) the corner domain 6 is

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. Figure 25-1 Three-dimensional model of RyR topology by cryo-electron microscopy. The cryo-EM data deposited at the EBI Macromolecular Structure Database, EMD-entry: 1,275 (Ludtke et al., 2005), was used to produce these figures for illustration purposes (http://www.ebi.ac.uk/msd-srv/emsearch/form). The electron-density map was converted to the CCP4 format and viewed using Pymol. The figure shows representative views of the electron-density (a) from the top (i.e., cytosolic side), (b and d) from the side or lateral views, (c) from the bottom (i.e., luminal side). The numbers indicate the domains reported by the Wagenknecht lab, or the more descriptive nomenclature (i.e., clamp, handle domain) used by the Chiu lab. Approximate dimensions estimated from the electron-density map are also indicated

connected to domain 4 which in turn is attached to another ‘‘handle’’ or domain 3, (3) a connection between domain 5 and domain 2. The corner domains appear to be formed by two distinct structures, a tubular structure that wraps around the corners of the square prism (domains 8a–8–7) and a smaller projection extending by the side of the square towards the corner (domains 9–10). Domain 10 projects outwards and appears to be kinked. A rhomboid structure is defined by domains 2–4–5–6, and appears to be connected to most of the other RyR1 domains. In addition, four columns 14 A˚ in height appear to constitute a bridge between the cytoplasmic region and the TM domain (Samso et al., 2005).

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3.2 Transmembrane Assembly The TM region appears to also have the overall shape of a square that is rotated by about 40 degrees from the square outlined by the cytoplasmic assembly (> Figure 25-1). When viewed on its side, the TM region appears to be tapered; the diameter of the end connected to the cytoplasmic domain being larger (120 A˚) than the distal end that protrudes into the lumen. The TM assembly corresponding to the mushroom ‘‘stalk’’ is approximately 70 A˚ in length, more than sufficient to traverse the membrane bilayer since lipid bilayers have an approximate thickness of 30–40 A˚ (Mitra et al., 2004). The shape of the TM region varies slightly, depending on the functional state of the channel when the RyR is analyzed by cryo-EM. For example, if the receptor is treated with ryanodine, which locks the RyR into a modified open state, the TM region electron density map appears to be better defined (Orlova et al., 1996) than when viewing this region without ryanodine bound (Radermacher, 1994). It is difficult to visualize the TM helices using any dataset at a resolution of around 10 A˚. At 3 A˚ resolution one can begin to view molecular boundaries of amino acids and at 1.8 A˚ the amino acid side chains become apparent in electron density maps. However, from their cryo-EM data the Wagenknecht and Chiu groups have been able to identify regions of high density that form rod-like structures, hence their designation as the putative helices that comprise the TM portion of RyR. The cryo-EM electron density implies that two central rod-like structures lie in each subunit, proximal to the fourfold axis of the channel (helix 1 and 2; > Figure 25-2). For helix 1, each of the pore-lining helices is approximately 45 A˚ long which would be accounted for by 30 amino acids. The lower half of each helix appears to be kinked and bends away from the fourfold central axis of the channel (Ludtke et al., 2005). Thus, the luminal entrance of the RyR channel has a funnel-like structure formed by the helices, with an apparent diameter of 30 A˚. The helices that form the funnel, taper towards the cytoplasm, where it has a diameter of 15 A˚. The electron density corresponding to helix 2 appears to  be 22 A˚ long, and is tilted at an angle of 50 while pointing towards the channel’s fourfold axis. It forms ˚ an opening of approximately 7 A with the corresponding helices from the other three subunits (Ludtke et al., 2005).

4

RyR Domain Organization

Analysis of the RyR amino acid sequence reveals a multi-domain protein organization, shown schematically in > Figure 25-3. Since some of these structurally conserved domains have been solved experimentally in different proteins, it is reasonable to assume that the folding of these regions of homology within the RyR would adopt similar folding patterns. When possible, the locations of such domains are related to the cryoEM structural information that is currently available.

4.1 MIR Domain The protein mannosyltransferase, IP3R and RyR (MIR) domain, as shown in > Figure 25-4, elucidated in the crystal structure of the N-terminal fragment of the mouse IP3 receptor (IP3R) adopts a beta-trefoil structure (Bosanac et al., 2002). Within the RyR molecule, these domains are thought to represent discrete structural entities rather than tandem repeats within the sequence. The importance of this domain can be inferred from the fact that the minimal region of the IP3R that binds to IP3 comprises the third and fourth MIR domain and the N-terminal portion of the first ‘‘RyR and IP3-homology’’ (RIH) domain (> Figure 25-4) (Yoshikawa et al., 1996). Thus the IP3 binding site is not thought to be confined to a single domain region but is rather at an interface between domains. However, there is no evidence that the RyR molecule binds to IP3 and currently, the significance of the MIR domain in RyR is not well understood.

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. Figure 25-2 Transmembrane features of the RyR. A wire-frame representation of the RyR electron-density map (EMDentry:1,275; see > Figure 25-1 for details) showing a slab-view (a) illustrating where the electron density for rod-like structures are prominent and which may represent the pore-lining helices. The approximate location of residues G4934, D4938, D4945 is indicated; these may represent the ‘‘gating-hinge’’ found in other ion channels, as discussed in section 3.4 (Jiang et al., 2002). The putative arrangement of transmembrane helices from one subunit of the RyR tetramer is shown (b), as inferred from combination of hydropathicity profiles (Zorzato et al., 1990) and site-directed antibody approaches (Grunwald Meissner, 1995). The 10 TM nomenclature is used (Zorzato et al., 1990). M9 is shown representing the selectivity filter and M7 is shown as a tight transmembrane hairpin. Amino-acid sequence alignment (c), showing the high degree of sequence identity in the presumed selectivity filter and pore helix region of the three RyR and IP3 receptor isoforms (IP3R1, IP3R2, IP3R3). The lower sequence is from the bacterial KcsA channel. The selectivity filter and putative ‘‘gating-hinge’’ is also conserved in the mammalian shaker Kv1.2 potassium channel (d) (PDB ID: 2A79). The arrangement of helices highlighting the position of the selectivity filter is shown from three different viewing angles; the left side shows a lateral view, right side shows the pore from cytosolic (top) or luminal side (bottom)

4.2 SPRY Domain The sequence repeats found in the dual-specificity kinase ‘‘splA and ryanodine receptor,’’ lends its name to the SPRY domain which is also found in the RyR molecule but not in the IP3R protein (see > Figure 25-3). This domain is found in triplicate in the N-terminal half, between RyR residues 670 and 1,561. SPRY domains are widely regarded to be protein-protein interaction domains that are involved in a number of biological functions such as innate retroviral restriction and in cytokine signaling. The SPRY domain is comprised of a single-domain anti-parallel b sandwich fold with one or two a helices (see > Figure 25-4).

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. Figure 25-3 Bioinformatics analysis of domain distribution in IP3 and ryanodine receptors. A schematic diagram to illustrate the various predicted domains and transmembrane segments within the human RyR1, RyR2 and IP3R1 sequences derived from bioinformatics analysis using the SMART database

The protein-protein interaction function is facilitated by a conformationally rigid and structurally defined binding pocket as determined in the SPRY domain of the suppressor of cytokine signaling (SOCS-box) protein (Woo et al., 2006).

4.3 EF Hand Domains and Ca2+ Binding Activation of the cardiac RyR2 isoform is triggered via the influx of Ca2+ ions through a voltage-activated plasma membrane Ca2+ channel, the dihydropyridine receptor (DHPR). In the skeletal muscle, the DHPR molecule is closely associated with the RyR1 isoform and a DHPR conformational change directly triggers the activation of RyR1. Cytoplasmic Ca2+ ions can act as an agonist for both isoforms and the released Ca2+ further activates adjacent RyR molecules in a phenomenon known as Ca2+ induced Ca2+ release (CICR). Based on the modulation of RyR channel function by variations in both cytosolic and luminal Ca2+, the existence of a number of functionally distinct Ca2+ binding sites on RyR has been postulated. The RyR is considered to contain sites where Ca2+ binding causes either activation of the channel (A sites) or sites where Ca2+ binding causes inactivation (I sites) (Laver, 2007). Topologically, these Ca2+ binding regions appear to be localized on the solvent-accessible surface of the RyR as shown by antibody binding studies (Du et al., 1998). Two EF hand domains are found towards the Cterminal portion of the RyR molecule (see > Figure 25-3). A 30 amino acid stretch corresponding to the calmodulin binding region of the RyR (residues 3,614–3,643 in rabbit RyR1) is able to bind to these EF hand motifs (amino acids 4,064–4,210 in rabbit RyR1) and alters the Ca2+ dependence of ryanodine binding (Xiong et al., 2006). Calmodulin (CaM), a 16.7 kDa naturally occurring protein containing EF- hand domains, has been shown to bind within a cytosolic cleft of the RyR (Wagenknecht et al., 1997).

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. Figure 25-4 Structure of discrete protein domains that are predicted in RyR. The crystal structure of protein domains that are predicted to be in the RyR, such as the MIR, SPRY, RIH and EF-hand domains are illustrated. The sequence alignment shows part of the IP3 binding site at the N-terminal end of the IP3R1, for which a crystal structure has been obtained (1N4K), aligned with a corresponding region near the N-terminus of the RyR2, revealing a similar helical folding pattern

CaM binding to the cytoplasmic domain of RyR1 affects SR Ca2+ release both in the presence or absence of Ca2+ ions (Meissner, 1986). The open probability of RyR1 is increased when Ca2+-free calmodulin (apoCaM) interacts with the channel, whereas Ca2+-CaM is inhibitory to channel opening (Buratti et al., 1995; Ikemoto et al., 1995; Tripathy et al., 1998). The CaM-binding region of RyR1 appears to correlate with domain ‘‘7’’ (Wagenknecht et al., 1997), which corresponds to the region encompassing amino acids 3,614–3,643, since addition of this synthetic peptide to SR vesicles modulates RyR1 function (Zhu et al., 2004; Xiong et al., 2006). The sequence encompassed between amino acids 3,614–3,643 is well conserved across all vertebrate RyRs and point mutations within this region affects CaM regulation of channel function in all RyR isoforms (Yamaguchi et al., 2001; O’Connell et al., 2002;

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Yamaguchi et al., 2003; Yamaguchi et al., 2005). As noted above, 3,614–3,643 has been shown to bind to the calmodulin-like EF-hand region 4,064–4,210 and this region awaits confirmation of the characteristic CaM folding pattern by crystallography. The implication of this finding is that the EF-hand motifs in region 4,064– 4,210 could compete with calmodulin for interaction with 3,614–3,643 as part of the complex Ca2+-sensitive mechanism of CaM-mediated channel regulation (Xiong et al., 2006). A recent crystallographic study has shown that CaM could be co-crystallized with a peptide fragment of RyR 3,532–3,557 (Maximciuc et al., 2006). The results of this study along with FRET measurements imply that one of the lobes of CaM may form contacts with region 3,532–3,557 of RyR while the other lobe may be free to interact with another part of the RyR (Maximciuc et al., 2006). The collective data from various laboratories provide strong evidence that both the regions 3,614–3,643 and 4,064–4,210 are solvent exposed and may be good candidates for recombinant expression and subsequent structural analysis. The structure and function of CaM depends on whether Ca2+ is bound to each of its two modules (Finn et al., 1995; Zhang et al., 1995). The two distinct forms of CaM exert opposite effects on the RyR. Both apoCaM and Ca2+-CaM bind to the cytoplasmic assembly with a stoichiometry of one CaM per RyR1 subunit (Moore et al., 1999), however, the positioning of CaM binding to RyR differs depending on the Ca2+ concentration. Ca2+-CaM binds within the crevice formed by domains 3 and 5/6 (Wagenknecht et al., 1994; Wagenknecht et al., 1997), while apo-CaM binds to the external part of domain 3 (Samso Wagenknecht, 2002). There appears to be a significant distance of 33  5 A˚, separating the binding of the 1999). Examination of the RyR sequence shows a separation of only 5 residues which is somewhat surprising when one considers the 33  5 A˚ separation observed by cryo-EM, which exemplifies the need to understand the 3D structural folding of the CaM binding domain of RyR. It may be that an evolutionary advantage exists with having two CaM binding sites in close proximity to each other, as it would enable a faster response if CaM interaction with RyR can switch rapidly between successive Ca2+ cycles without requiring complete dissociation from the RyR.

4.4 Transmembrane Domains The TM domains are often regarded as the most important domain in an ion-channel, since they define the pore segment and anchor the protein in the lipid bilayer. There is also evidence that TM domains contain the determinants for the oligomerization state of a membrane receptor (Galvan et al., 1999). For the RyR, hydropathy profiles have predicted between 4 and 12 membrane-spanning sequences near the C-terminal end (Takeshima et al., 1989; Zorzato et al., 1990; Brandt et al., 1992; Tunwell et al., 1996). However, hydropathy profiles are interpreted with caution, since some peaks in a hydrophobicity profile can also represent non-polar pockets within soluble domains that are not regions embedded in a lipid bilayer. Improved algorithms for prediction of TM helices using hidden Markov models, such as the TMHMM server (Krogh et al., 2001) and the Phobius server (Kall et al., 2004), are currently available. The nomenclature for the 12 transmembrane (12TM) model of RyR is still in use (Zorzato et al., 1990) and a site-directed antibody approach has indicated the ninth TM segment (M9) in the 12TM model as a luminal loop (Grunwald Meissner, 1995). It is suggested that the path of the Ca2+ ion traversing the channel is lined by the TM helices M9 and M10 (Zorzato et al., 1990), and relating this to the RyR cryo-EM structure, M10 is most likely represented by helix 1 as schematically shown in > Figure 25-2 (Du et al., 2004). The sequence GXXXA has been proposed to form a ‘‘gating’’ hinge in the inner, pore-lining helix within potassium channels (Jiang et al., 2002). Molecular modeling studies carried out by the Williams group have suggested that a similar sequence is found in RyR2 between residues of G4934 and A4939 (Welch et al., 2004). The Chiu group have proposed that helix 1 is formed between I4918 and E4948 with the kink being represented by G4934. The P-loop (pore-loop) associated helix is likely to correspond with M9 or helix 2, entering the channel from the luminal side of the membrane (Zhao et al., 1999; Gao et al., 2000). The ‘‘GGGIGD’’ (amino acids 4,894–4,899) sequence in the luminal loop between M9 and M10 (> Figure 25-2) is analogous to the selectivity filter (TVGYGD) in potassium channels (Jiang et al., 2002). Mutagenesis of residues in this

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region of both RyR1 and RyR2 alters channel conductance, thus offering evidence to support the assignment of this segment of amino acids as the selectivity filter (Zhao et al., 1999; Gao et al., 2000; Du et al., 2001; Chen et al., 2002).

4.5 FKBP Binding Domain The drug tacrolimus, or FK506, was initially isolated from Streptomyces and its potency was reported initially to exceed that of cyclosporine, an immunosuppressant used in transplant surgeries (Kino et al., 1987). A novel protein that bound to FK506, distinct from cyclophilin the natural target protein of cyclosporine, was identified and named FK506 binding protein (FKBP) (Siekierka et al., 1989a, b). FKBP was shown to possess a ‘‘cis-trans isomerase’’ activity (Fischer et al., 1989; Harding et al., 1989). Following the discovery of FK506 and its ability to prevent graft-versus-host rejection in organ transplantation, a flurry of research activity led to the identification of RyR as a binding partner of FKBP (Jayaraman et al., 1992). The amino-acid sequence alignment of two human FKBP isoforms, FKBP12 and FKBP12.6, is shown in > Figure 25-5. There is 78% sequence identity between these isoforms; FKBP12 binds predominantly to RyR1 whereas FKBP12.6 binds to RyR2. FKBP 38, another related isoform is also illustrated in the sequence alignment and plays an indirect role in cardiopathophysiology. FKBP38 interacts directly with RheB, which in a GTP-dependent manner prevents its association with mTOR, a protein that is implicated in the pathways for cardiac hypertrophy, chronic pressure-overload and cardiac fibrosis. Both FKBP12 and FKBP12.6 have a high affinity binding interaction with the RyR and this feature has enabled affinity purification of RyR1 using immobilized FKBP12 (Xin et al., 1995; Bultynck et al., 2001a). The dissociation constant (Kd) of FKBP12 for the ‘‘open’’ RyR channel is 1nM, whereas the Kd of FKBP for the ‘‘closed’’ RyR channel is 0.1pM, indicating a very strong association between FKBP12 and RyR1 in maintaining the ‘‘closed’’ state of the channel (Jones et al., 2005). Cryo-EM studies have localized FKBP binding to a region adjacent to domain 9, at the corner of the RyR cytoplasmic domain (> Figure 25-5) (Sharma et al., 2006). The fitting of FKBP into the electron density map of the RyR-FKBP complex suggests that Q31 and N32 of FKBP interact with the exposed surface of RyR. Rapamycin, like FK506, is a natural target ligand for FKBP12/12.6 that is typically used to dissociate FKBP from the RyR. In this cryo-EM study, the FKBP pocket that normally binds rapamycin is occluded when FKBP binds RyR, raising the issue of how rapamycin can access its interaction site to cause dissociation of the FKBP-RyR complex. The cryo-EM evidence suggests that a phenylalanine residue within the FKBP hydrophobic pocket that binds rapamycin and FK506 is 10 A˚ from the nearest RyR surface residues (Sharma et al., 2006). Thus, there is a small gap for rapamycin to fit in between the FKBP and RyR that causes the dissociation of FKBP. Another study of the FKBP12-RyR1 complex has shown that domains 3 and 9 are also involved in the interaction with FKBP (Samso et al., 2006). This study advocated a rigid-body docking method as opposed to the computationally rigorous flexible-ligand docking approach and suggested that a FKBP loop comprising amino acids 82–95 interact with domain 3 (see > Figure 25-5) and that residues 87–90 are pointed towards the RyR (Samso et al., 2006). As FKBP is implicated in the regulation of channel functional state (open or closed), there has been considerable interest in mapping its binding site(s) on the RyR sequence, but the precise FKBP12 binding domain of RyR remains controversial. FKBP binding has been identified within the RyR N-terminal half (Lee et al., 2004) and GST-FKBP affinity pull-down assays have shown interaction with the N-terminal 1,855 residues (Masumiya et al., 2003; Xiao et al., 2004). In contrast, binding regions identified by yeast two-hybrid analysis have implicated a short central region incorporating amino acids 2,407–2,520 of RyR1 (Cameron et al., 1997) and 2,361–2,496 of RyR2 (Marx et al., 2000). Site-directed mutagenesis of RyR1 (Gaburjakova et al., 2001) and RyR3 (Bultynck et al., 2001b; Van Acker et al., 2004) have also implicated the RyR central region in FKBP binding. These studies identified V2461 in RyR1 (and the corresponding V2322 in RyR3) as an important residue for FKBP binding. Evidence from our laboratory has implicated binding sites in the C-terminal half of the molecule (Zissimopoulos Lai, 2005). In addition to this, it has been suggested that a C-terminal residue in RyR2, P4595 (numbering for human RyR2) may interact with FKBP (Jayaraman et al., 1992; Ahern et al., 1994; Brillantes et al., 1994). Proline residues are

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. Figure 25-5 FKBP and RyR interaction. The approximate location of FKBP binding to the corner of RyR at domain 9 is shown (a) (Sharma et al., 2006). Amino-acid sequence alignment for human FKBP12, FKBP 12.6 and FKBP 38 (b) is presented. On the right is a ribbon representation of FKBP12 (PDB:2FAP), indicating a putative region that interacts with RyR. Residue I56 of the crystal structure forms a polar contact with an oxygen atom in rapamycin, with F46 also implicated in a hydrophobic interaction to maintain rapamycin binding to FKBP12. The potential interaction between RyR1 and FKBP12 is shown (c), based on theoretical docking of the ribbon-representation of FKBP12 and RyR1 (Sharma et al., 2006)

of particular significance since FKBP is a cis-trans proline isomerase and proline has an unusual conformationally-restrained peptide bond due to its cyclic side chain. The ‘‘trans-peptide-bond’’ conformation is usually preferred in most amino acids within a protein sequence due to considerations of steric hindrance; however the unusual structure of proline allows the cis form to be stable as well. Hence, both cis and trans isomers are found in proteins. These discrepancies in the determination of FKBP binding sites may be due to intrinsic limitations of the experiment, since most of the assays in site-mapping studies have used some form of solid-support matrix (e.g., GST pull-down assays) in which the FKBP would be held in an undetermined fixed conformation. Further studies using methods such as isothermal titration calorimetry (ITC) (Velazquez-Campoy et al., 2004) may help resolve these controversies, since proteins in an ITC chamber are unrestricted, allowing interactions to occur in natural conformations.

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4.6 RyR Phosphorylation Reversible protein phosphorylation and dephosphorylation are frequent events in biology which determine receptor or enzyme activation/inactivation by effecting a protein conformational change. Phosphorylatable amino acids in eukaryotes are serine, threonine and tyrosine, while in prokaryotic proteins, histidines, arginines and lysines are also phosphorylated (Cozzone, 1988). The human genome has at least 518 protein kinases (Manning et al., 2002), with most exhibiting significant structural homology hence their grouping into a kinase superfamily comprising two groups; serine/threonine kinases and tyrosine kinases, usually known as ‘‘conventional’’ protein kinases (CPKs). CPKs phosphorylate residues that are located in loops, turns or irregular structures (Pinna Ruzzene, 1996), whereas ‘‘alpha kinases’’ e.g., elongation factor-2 kinase phosphorylate residues that are located in alpha helices (Ryazanov et al., 1999). Thus, resolving the kinase involved in a phosphorylation event may inform on potential tertiary folding of the substrate. Purified RyR phosphorylated in vitro by cAMP-dependent protein kinase showed a 30% increase in ryanodine binding (Takasago et al., 1989), suggesting this modification favors the open channel state. RyR is also phosphorylated by Ca2+/calmodulin-dependent kinase (CaMKII) and protein kinase A (PKA) (Seiler et al., 1984; Takasago et al., 1989; Frey et al., 2000), both CPKs, suggesting that the RyR phosphorylation site may adopt a solvent-accessible loop or turn. Phosphorylation of the RyR2 residues S2031, S2808 and S2814 have been demonstrated and additional sites were proposed using bioinformatics analysis (George, 2008). The phosphorylation of only one/a few residues within the huge oligomeric RyR can significantly alter channel regulation. Several studies have shown that RyR phosphorylation affects the binding of accessory proteins, such as FKBP. Stoichiometric labeling (termed ‘‘hyperphosphorylation’’) of a single residue, S2808, has been proposed to directly disrupt the binding of FKBP12.6 to RyR2, thus favoring a ‘‘leaky’’ channel state and triggering an arrhythmic event that can lead to sudden death (Marx et al., 2000; Wehrens et al., 2003). These results imply that the S2808 residue is involved in binding to FKBP12.6. However, recent cryo-EM studies of a GFP tag inserted after Y2801 in RyR2 indicated that this location is at a distance of 105–120 A˚ from the previously mapped FKBP site (Meng et al., 2007). Due to the lower resolution of cryoEM studies this data should be interpreted with caution, and because it is feasible that GFP tag insertion into the RyR results in a structural perturbation that is transferred to a distal region of the complex. The RyR binding sites for FKBP, CaM, putative phosphorylation sites, locations of putative TM helices and other features as indicated by cryo-EM studies and in vitro experiments are summarized in the schematic diagram shown in > Figure 25-6.

4.7 DHPR-RyR Interaction Domain DHPRs reside in the transverse tubules closely apposed to RyR at the SR terminal cisternae regions, thus facilitating the E-C coupling mechanism in skeletal muscle whereby a voltage-induced DHPR conformational change is instantly communicated to the directly coupled RyR1. This signal switches the RyR1 to an open state thereby enabling Ca2+ release from the SR to elicit muscle contraction (Schneider Chandler, 1973). Imperatoxin A (IpTxa), a high-affinitypeptide mimetic of a cluster of basic residues at the DHPR Nterminus (residues 666–791; Gurrola et al., 1999) has been used to elucidate the DHPR interaction site on RyR (Samso et al., 1999). Binding of IpTxa enhances the interaction of RyR with DHPR (el-Hayek et al., 1995) and encourages long-lived subconductance states (Tripathy et al., 1998; Gurrola et al., 1999). CryoEM analysis of streptavidin-labeled IpTxa incubated with RyR1 revealed that IpTxa binding was localized to the crevice formed by domains 3 and 7/8 of the cytoplasmic assembly (Samso et al., 1999). An allosteric Ca2+ release mechanism is likely, since the distance of the IpTxa binding site from the channel pore is significant (110 A˚) and the distance between neighboring DHPRs within a tetrad approximates that observed between the centers of mass of neighboring IpTxa (150 A˚) (Block et al., 1988). The binding site for IpTxa is very close to that for Ca2+-CaM according to cryo-EM studies. If IpTxa is indeed a true representation of the DHPR II-III loop:RyR1 interaction, then CaM may play a direct role in E-C coupling protein interactions.

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. Figure 25-6 Functional domains identified within the RyR. The location of various RyR functional domains that have been empirically determined are shown in (a). The RyR2 isoform is shown here, although the pattern is similar for RyR1. The schematic is drawn approximately to scale using VectorNTI (Informax). The topology defined by cryoEM studies for interaction with various RyR associated proteins, as well as for the three RyR divergent regions involving use of GFP insertion is shown in (b)

4.8 RyR Binding to Accessory Proteins A large number of additional accessory proteins are known to bind to the RyR including calmodulin, calsequestrin, triadin, junctin, sorcin, homer, and various toxins (Bers, 2004). The identification of the RyR interaction sites for these accessory proteins and their role in influencing RyR protein folding and function

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is a vital area of continuing investigation. The physiological effect of modifications to these RyR accessory proteins is of extreme importance. For example, transgenic mice carrying the human cardiac calsequestrin D307H missense mutation or the calsequestrin-null mutation showed structurally normal hearts but developed stress-induced ventricular arrhythmias (Song et al., 2007). These mice exhibited cardiac hypertrophy and reduced contractile function with age progression, and also displayed a reduction in calsequestrin gene expression combined with a compensatory elevation in calreticulin and RyR2 expression, which could explain the premature Ca2+ release observed in cardiac myocytes (Song et al., 2007). Natrin, a 25 kDa snake venom toxin from Naja naja atra, is a member of the cysteine-rich secretory protein (CRISP) family (Wang et al., 2004; Wang et al., 2006). CRISP family members are predominantly found in the mammalian male reproductive tract as well as in the venom of reptiles, and many have been shown to have a direct effect on ion channel function. The CRISP Tpx-1 domain has been shown to regulate RyR Ca2+ signaling (Gibbs et al., 2006).

5

Comparison of RyR Isoforms

Bioinformatics analysis (ClustalW) of the three RyR isoforms indicate a high degree of sequence identity (66–70%), however this has also identified three discrete non-homologous regions, known as the ‘‘divergent regions’’ 1, 2, and 3 (DR1, DR2, and DR3) (Sorrentino Volpe, 1993). DR1 is the largest divergent region comprising of 360 residues (RyR1 4,254–4,631; RyR2 residues 4,210–4,562). The two other divergent regions are much smaller, whereby DR2 region includes 50 residues (RyR1 1,342–1,403; RyR2 1,353–1,397) and the DR3 region has 45 residues (RyR1 1,872–1,923; RyR2 1,852–1,890) (> Figure 25-6). These specific regions of sequence divergence have been the focus of a number of structural and functional studies, as it is thought that they are likely to play a major role in the distinct physiological properties of the individual RyR isoforms. Cryo-EM studies of RyR2 containing a GFP insertion after residue T1366 have suggested that this increased GFP mass placed within the DR2 region produces a corresponding increase in domain 6 of the RyR topology model (> Figure 25-6) (Liu et al., 2004). Similarly, using a GFP insertion after RyR2 residue T1874 in the DR3 region the increase in mass was mapped to domain 9, in the clamp region adjacent to a proposed FKBP-binding site (Zhang et al., 2003). The GFP-tag inserted after RyR2 residue D4365 enabled the proposed location of DR1 to domain 3, or the handle region of RyR (Liu et al., 2002). The proposed location of the divergent regions and other salient features of the domains within the RyR topology model are shown in > Figure 25-6. To date, of the three isoforms, only RyR1 has been reconstructed by cryo-EM to a higher degree of structural resolution (Serysheva et al., 2008). The RyR2 and RyR3 isoform topology currently remain at a lower resolution. As may be anticipated from the 70% sequence identity between the three mammalian RyR isoforms, the 3D reconstructions of RyR2 and RyR3 are nearly identical to that of RyR1. However, there is one significant difference in the topology of RyR3 compared with that of RyR1 and 2, corresponding to domain 6 of the clamp region, where the absence of a mass of 10 kDa is observed in RyR3, but which is present in both RyR1 and 2. This mass difference appears to be due to the RyR3 sequence length being 100 residues shorter, specifically in the region corresponding to residues 1,303–1,406 of the other two isoforms.

6

Open and Closed States of RyR

As with any channel protein, it is interesting to note the differences that exist when the channel is in the open or closed state. In the cryo-EM models for RyR, the presumed open or closed channel conformation is induced with specific buffer conditions (Sharma et al., 2006). In the ‘‘open’’ state, the clamp domains at the four corners of the RyR cytoplasmic region appear to adopt a more open conformation. The TM regions also show a structural perturbation that apparently results in forming a central ‘‘hole’’ in the open state that is not observed in the closed state. The channel opening mechanism has been compared to the openingclosing of the iris in a camera (Serysheva et al., 1999). Others have noted no appreciable differences between

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the ‘‘open’’ and ‘‘closed’’ states for RyR1 and RyR3 (Sharma et al., 2000). However, an interesting aspect of channel opening and closing is the observable change in the clamp regions that are positioned 100 A˚ from the changes that occur in the TM region. How the conformational change in a peripheral cytoplasmic domain is related to the presumed functional and observed structural change in the TM region during channel opening and closing remains to be clarified. A plausible explanation could involve the clamps mediating direct interactions between DHPR and RyR1, a mechanism that appears to be an essential component of E-C coupling in the skeletal muscle (Paolini et al., 2004a, b; Sheridan et al., 2006). Hence, conformational changes in the clamp structure accompanying signal communication from DHPR to the RyR would be transmitted to the TM region and result in channel gating.

7

Interaction Between RyRs

The interaction between RyRs also deserves particular attention because of the key role it may play in the phenomenon of ‘‘Ca2+ sparks.’’ These are large Ca2+ release events that seem to be mediated by the coordinated opening of groups of interconnected RyR channels and constitute the fundamental physiological Ca2+ signaling process in muscle cell contraction. In the SR membrane, RyRs are packed into regular lattices that exhibit a unique ‘‘checkerboard’’ pattern, whereby individual square-shaped RyR oligomers are in proximity at each of the four corners (Block et al., 1988; Saito et al., 1988). This regular array of RyR particles raises two central questions. Why do the RyRs organize into such a unique pattern, and which particular RyR domain(s) is involved in the interaction to form the 2D array? From a crystallographic perspective, the closest packing arrangement for RyRs would be a ‘‘side-by-side’’ pattern, as the four identical receptor subunits arrange symmetrically around a central fourfold axis. Then why do the RyRs organize into lower-density checkerboard arrays and not pack into high-density side-by-side arrays? To facilitate the unique RyR organization in SR membranes, we hypothesized a specific RyR-RyR interlocking of oligomers exists, so that each RyR within native arrays is coupled through a specific physical link to a neighboring RyR, to establish the stable checkerboard lattice (Yin et al., 2008). Electron microscopy and image analysis of checkerboard arrays revealed that: (1) adjacent RyRs are indeed interlocked through a physical link, and (2) the inter-oligomeric interaction occurs via the distal edge of domain 6, at the corner of each clamp region (> Figure 25-7; Yin et al., 2005a). RyR association into native ‘‘checkerboard’’ arrays is ionic strength-dependent, suggesting a role for an electrostatic component in the protein-protein interaction between RyR neighbors (Yin & Lai, 2000, Yin et al., 2005a, b). Intriguingly, the RyR DR2 region also maps to the distal edge of domain 6 (> Figure 25-6), which corresponds to the interaction domain linking adjacent RyRs within the checkerboard array (> Figure 25-7; Yin et al., 2005a; Yin et al., 2008). Notably, this RyR region comprises alternating sections of positively-charged (1383-KNKKRGFLFKAKK-1395 in RyR1, 1383-RLKQRFLLRRTK-1394 in RyR2; +ve charge residues in blue italics) and negatively-charged (1320EDEARAAEPDPDYE-1333 in RyR1, 1333-DLEDYDADSDFE-1344 in RyR2; ve charge residues in red italics) amino acids (Otsu et al., 1990; Zorzato et al., 1990). The specific distribution of alternating charged segments within this DR2 region of RyR1 and RyR2 suggests that these residues may play a role in mediating the physical coupling occurring between these RyR isoforms (Yin et al., 2008). Interestingly, absence of this pattern of charged amino acids in RyR3 would infer that 2D array formation may not be an intrinsic feature of this isoform.

8

Perspectives

Despite the elegant topological data obtained so far by various laboratories using advanced cryo-EM techniques, there remains a compelling desire for the more detailed structure of RyR at atomic resolution. The enormous size of the RyR monomer and oligomer is certainly a great challenge for the application of X-ray crystallography which requires as a starting point, homogenous preparations of RyR in high milligram quantities. Without such detailed atomic structures, we can only speculate as to the nature of the selectivity filter, the residues that form the pore helices and how minute changes at the TM domains are

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. Figure 25-7 Physical coupling between RyRs and with DHPR. The physical coupling between RyRs and DHPR in skeletal muscle is illustrated in (a), based on freeze-fracture studies and 3D reconstructions of these two Ca2+ channels generated by cryo-EM and single particle analysis. The RyR array (grey) is overlaid by DHPR particles (dark grey, grouped as tetrads). The square box outlines one RyR1 tetramer and its interaction with a DHPR tetrad. Note ˚ . Reproduced from that the tetrads interact with every second RyR particle (Yin et al., 2008). Scale bar = 100 A Serysheva (2004). The projection map in (b) shows the ‘‘checkerboard-like’’ pattern arrangement of the RyR1 array. The square box outlines one RyR1 molecule which is physically linked through the distal edge of domain 6 to four neighboring RyR molecules. A model of the physical coupling between RyRs in the native checkerboard array is shown in (c), constructed according to domain assignment, which matches the in situ lattice constants (a = b = 31 nm, Block et al., 1988; Saito et al., 1988). The RyR1 3D structure of Radermacher et al. (1994; > Figure 25-1a) was used as a molecular model in building the array

related to simultaneous perturbations within the cytoplasmic domain. Whereas cryo-EM and singleparticle image processing techniques can in principle achieve atomic resolution, and recent advances in viruses with a high degree of symmetry are indeed approaching atomic resolution (3.8 A˚; ZH Zhou, UCLA, personal communication), this has not yet been achieved for complexes with lower symmetry. To date, the highest resolution reported using these techniques for complexes that lack high symmetry is 7.5 A˚, which was obtained for a 50S ribosomal subunit (Matadeen et al., 1999). Since the first 3D reconstruction of RyR1 was reported in 1994 (Radermacher et al., 1994), the resolution of reported reconstructions has improved from 30 to 10 A˚ (Ludtke et al., 2005; Samso et al., 2005; Serysheva et al., 2008). In attempting to achieve higher resolution structure information for the RyR, various laboratories are continuing to make extensive efforts and there has been some promising progress made, including the preparation of large 2D crystals of RyR (5 mm) that diffract to 20 A˚ with negative stain (Yin et al., 2005a, b). Small 3D crystals of purified

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native RyR protein have also been obtained with defined edges (George et al., 2005). Currently, we are continuing our efforts to refine the crystallization conditions to produce improved 2D and 3D crystals amenable to cryo-electron crystallography and X-ray crystallography.

Acknowledgements We Wish to thank the British Heart Foundation, European Commision, Medical Research Council and Wellcome Trust for supporting our research.

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Signal Molecules and Calcium

N. Damann . D. D’hoedt . B. Nilius

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 490

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The TRP Subfamilies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 492 TRPC . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 492 TRPV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 493 TRPA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 496 TRPM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 497 TRPML . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 499

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Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 500

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2009 Springer Science+Business Media, LLC.

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Abstract: The 28 mammalian members of the superfamily of transient receptor potential (TRP) channels are cation channels, mostly permeable to both monovalent and divalent cations, and can be subdivided into six main subfamilies: the TRPC (canonical), TRPV (vanilloid), TRPM (melastatin), TRPP (polycystin), TRPML (mucolipin), and the TRPA (ankyrin) groups. TRP channels are widely expressed in a large number of different tissues and cell types and their biological roles appear to be equally diverse. In neurons of the peripheral and central nervous system, TRP channels may substantially contribute to neuronal excitation, neurotransmitter release, and calcium homeostasis. Some TRPs are involved in sensory functions such as temperature and pain perception and may help to regulate body temperature or to avoid tissue-damaging stimuli like noxious temperatures or irritating chemicals. Other TRPs appear to be intimately connected to G-protein-coupled receptors, thereby linking chemical signals such as hormones and neurotransmitters to membrane excitability. TRPs may even be involved in regulation of neurite length and growth cone morphology. In this chapter, we provide an overview of the impact of TRP channels on cellular calciumdependent processes in mammalian neurons and identify several TRPs for which a causal pathogenic role might be anticipated. List of Abbreviations: ADPR, ADP-ribose; AITC, allyl isothiocyanate; DRG, dorsal root ganglion; HEK, human embryonic kidney; NCCa-ATP, nonselective Ca2+-activated channel; NMDA-R, NMDA receptors; OGD, oxygen–glucose deprivation; OVLT, organum vasculosum lamina terminalis; PAG, periaqueductal gray; PIP2, phosphatidylinositol bisphosphate; PLC, phospholipase C; PNS, peripheral nervous system; ROCE, receptor-operated calcium entry; TRP, transient receptor potential; VACCs, voltage-activated Ca2+ channels

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Introduction

Transient receptor potential (TRP) channels constitute a large and functionally versatile family of cation conducting channel proteins. The first TRP channel gene was discovered in Drosophila melanogaster (Montell and Rubin, 1989) in the analysis of a mutant fly whose photoreceptors failed to retain a sustained response to a long light stimulus (Cosens and Manning, 1969) (for a review, see Wang and Montell, 2007). In the following decade more than 50 TRP channels were identified with representative members in many species right through from yeast to human (for review, see Nilius and Voets, 2005; Pedersen et al., 2005; Voets et al., 2005; Nilius et al., 2007). In mammals, 28 TRP channels were found and classified according to homology into six subfamilies (Montell et al., 2002): TRPC (Canonical), TRPV (Vanilloid), TRPM (Melastatin), TRPA (Ankyrin), TRPML (Mucolipin), and TRPP (Polycystin). TRPs are expressed in numerous neuronal and nonneuronal tissues where they are involved in manifold physiological functions, ranging from vasorelaxation, pheromone sensory signaling, fertility, taste transduction, nociception, temperature sensation, and osmoregulation. We are at the very beginning to identify all the diverse physiological functions of this intriguing ion channel family and our knowledge about TRP channel expression and functioning in various tissues of mammals is limited but accumulating evidence suggests that TRP channels play prominent roles in the regulation of the intracellular calcium level in both excitable and nonexcitable cells. The molecular architecture of TRP channels is reminiscent of ligand- and voltage-gated channels (Yu and Catterall, 2004) and comprises six putative transmembrane segments (S1–S6), intracellular N- and C-termini, and a pore-forming reentrant loop between S5 and S6. The length of the cytosolic tails varies greatly between TRP channel subfamilies, as do their structural and functional domains (for detailed reviews, see Clapham, 2003; Vriens et al., 2004a; Owsianik et al., 2006). By analogy to other channels with a similar transmembrane structure, TRPs probably form a tetrameric quaternary structure (Hoenderop et al., 2003) with each subunit contributing to a shared selectivity filter and ion-conducting pore similar to that seen in K+ channels (Long et al., 2005; Owsianik et al., 2006). Increasing evidence suggests heteromultimeric channel assembly within one subfamily, creating a variety of different channels with unique properties as compared to homomers (Strubing et al., 2001; Smith et al., 2002).

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TRP channels contribute to changes in the cytosolic free Ca2+ concentration either by acting as Ca2+ entry pathways in the plasma membrane or via changes in membrane polarization, modulating the driving force for Ca2+ entry mediated by alternative pathways. All TRPs [except TRPM4 and TRPM5 (Launay et al., 2002; Prawitt et al., 2003)], are permeable for Ca2+ and show moderate Ca2+ selectivity [permeability ratio relative to Na+(PCa/PNa) between ~0.3 and 10], with TRPV5 and TRPV6 exhibiting highly Ca2+-selective channel pores (PCa/PNa > 100). TRP channels can function as cellular sensors being directly gated by endogenous or extracellular stimuli and some TRPs are activated following G-protein-coupled receptor stimulation and may promote receptor-operated calcium entry (ROCE) (Hofmann et al., 1999; Okada et al., 1999; Beck et al., 2006). In nonneuronal cells, TRPs are discussed to mediate ROCE either by being directly activated (e.g., TRPC3, TRPC6, and TRPC7) by the second messenger diacylglycerol (DAG) which is produced by receptor-activated phospholipase C (PLC) enzymes, or by functioning as Ca2+-selective store-operated channels (Ca2+-SOCs, also known as CRACs) upon Ca2+ release from internal Ca2+ stores, such as the golgi apparatus, the endoplasmic reticulum (ER), or specifically in muscle cells, the sarcoplasmic reticulum. Whether TRPs function as SOCs is still a matter of debate. Recently, a functional interaction between the stromal interaction molecules STIM1 and STIM2 and the SOC in Drosophila S2 and Jurkat cells, respectively, has been discovered. In brief, reduced expression of STIM1 disrupts SOC activation in response to depletion of intracellular Ca2+ stores (Liou et al., 2005; Roos et al., 2005; Zhang et al., 2005). STIM2 seems to act as an inhibitor of STIM1-dependent CRAC (Soboloff et al., 2006). It will be important to determine whether STIM proteins interact with TRPC members that have been advanced as putative SOCs. Other possible partners of SOC channels are the recently discovered four TM proteins Orai1 (also named CRACM1) (Feske et al., 2006; Vig et al., 2006). Overexpression of Orai1 and STIM1 induces a dramatic increase in the size of CRAC currents (Mercer et al., 2006; Peinelt et al., 2006; Soboloff et al., 2006). Very likely, Orai1 is the long-thought CRAC channel (Parekh, 2006a, b; Peinelt et al., 2006; Vig et al., 2006; Zhang et al., 2006). Several studies show now that TRPC1 associates with STIM1 (Huang et al., 2006; Lopez et al., 2006; Ong et al., 2007) and also with Orai1 (Ambudkar and Ong, 2007; Ong et al., 2007). TRPC1/STIM1 and Orai1 may form ternary complexes to contribute to a SOC channel (Ambudkar and Ong, 2007) and recent findings suggest that all TRPCs, except TRPC7, can function as SOCs when they interact with STIM1 (Yuan et al., 2007). In neurons, some TRP channels are implicated in processes including sensory neuron activation, release of neurotransmitters and neuropeptides, action in the spinal cord, and axon growth and guidance. Recent investigations on the mammalian peripheral nervous system (PNS) revealed expression of TRP channels in sensory neurons of the trigeminal and dorsal root nerves and give evidence for fundamental impact of TRP receptor activation on neuronal excitation and intracellular calcium levels in these cells. TRP channels constitute key receptor proteins for conversion of environmental stimuli into electrochemical signals (Julius and Basbaum, 2001) which are conveyed by sensory neurons to the central nervous system (CNS), where signals integration and interpretation results in appropriate reflexive and cognitive behavior. On a molecular level, some TRP channels, especially those expressed in sensory neurons, were found to have polymodal modes of activation, manifested by simultaneous sensitivity to chemicals, temperature, and intracellular molecular modulators. In addition, some TRP channels (e.g., TRPV1, TRPV2, TRPV4, TRPM3, TRPM8, TRPA1) exhibit a weak voltage-dependent gating behavior which is characterized by a shallow Boltzmann slope in channel conductance–voltage (G–V) relations (Nilius et al., 2005). The voltage dependence manifests as channel activation upon depolarization to positive transmembrane potentials (Brauchi et al., 2004; Voets et al., 2004a), a hallmark of neuronal and other excitable cells. TRP channels exhibit a small gating charge which might be an important evolutionary structural prerequisite for their gating diversity as small changes in Gibb’s energy induce large shifts in V1/2 of TRP channel activation. Thereby, TRPs act as molecular integrators of polymodal stimuli and are reciprocally modulated in their voltage sensitivity by temperature or ligand binding. In this chapter, we focus on TRP channels from the TRPC, TRPV, TRPA, TRPM, and TRPML subfamilies that were shown to be expressed in mammalian neurons. We illustrate their predicted functioning in the context of neuronal physiology and pathophysiology thereby considering recent advances that have shed insight into the molecular mechanisms of TRP-dependent regulation of intracellular Ca2+.

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Signal molecules and calcium

The TRP Subfamilies

2.1 TRPC TRPC channels constitute the closest mammalian homologs of Drosophila TRP (30–40% identity). In vertebrates, the TRPC subfamily consists of seven members (TRPC1-TRPC7) and is readily subdivided on the basis of sequence similarity and functional comparison into three subgroups: group 1 (TRPC2), group 2 (TRPC3, TRPC6, TRPC7), and group 3 (TRPC1, TRPC4, and TRPC5) (Montell et al., 2002). Coimmunoprecipitation for native TRPC channels in synaptosomes revealed that TRPC3, TRPC6 and TRPC7 can coassemble and that TRPC1, TRPC4, and TRPC5 can also coassemble with each other, whereas cross-assembling between the TRPC subgroups does not occur (Goel et al., 2002; Hofmann et al., 2002; Strubing et al., 2003) (for review, see Putney Putney, 2005). When compared to homomeric channels, heteromers might reveal modified biophysical properties as shown for TRPC1 when coexpressed with TRPC4 and TRPC5 (Strubing et al., 2001) and therefore, TRPCs constitute a large variety of channels with different qualities. All TRPC isoforms were shown to be expressed within the peripheral and central mammalian nervous system (Philipp et al., 1998; Strubing et al., 2001; Elg et al., 2007). As an exception, the TRPC2 gene is considered a pseudogene in humans where it encodes a nonfunctional truncated protein (Vannier et al., 1999) whereas in other mammalian species like rodents TRPC2 appears to be required for neuronal excitability in vomeronasal function (Liman et al., 1999; Stowers et al., 2002) (for review, see Zufall et al., 2005). In the hippocampus TRPC1, TRPC4, and TRPC5 were found at high levels in pyramidal cells of the CA1 region (Philipp et al., 1998; Chung et al., 2006) and especially TRPC5 could be detected in granule cells in the dentate gyrus (Chung et al., 2006). TRPC5 signals are also found in the mitral cells of the olfactory bulb and in lateral cerebellar nuclei (Philipp et al., 1998). Expression of TRPC1 transcripts and protein is more widespread, and often overlaps with that of TRPC4 and TRPC5 (Strubing et al., 2001). TRPC4 and TRPC5 can be activated by receptor stimulation and it was shown that Ca2+ has complex effects on these TRPC channels. Spontaneous currents through TRPC5 and receptor-mediated activation of TRPC4 and TRPC5 is dependent on intracellular Ca2+ and increasing extracellular Ca2+ (>5 mM) facilitates channel activity (Okada et al., 1998; Schaefer et al., 2000; Strubing et al., 2001; Jung et al., 2003). In the absence of extracellular Ca2+, activation was also found to be weak and subsequent addition of Ca2+ to the extracellular medium resulted in a large increase in current with a delay (Schaefer et al., 2000) suggesting that Ca2+ must first pass through the channel and accumulate within the cell. Also in inside-out patches, an increase in the Ca2+ concentration at the intracellular surface of the membrane increases currents through TRPC4 and TRPC5 in the absence of other activators (Schaefer et al., 2000). The action at the intracellular site is likely to be important during receptor-mediated channel activation where both Ca2+ release from intracellular stores and Ca2+ entry through the channel are likely to potentiate current activation. Uniquely among TRP channels, Gq/11 receptor-mediated activation of TRPC4 and TRPC5 (and of heteromers containing TRPC1) is potentiated by micromolar concentrations of the trivalent lanthanide cations La3+ of Gd3+ (Schaefer et al., 2000; Strubing et al., 2001), presumably by interacting with an extracellular Ca2+-binding site (Jung et al., 2003). The receptor-mediated activation mechanism of TRPC5 and TRPC4, together with their expression in central neurons, but not in glial cells, suggests that these channels may underlie currents and [Ca2+]i signals associated with the activation of G-protein-coupled muscarinic neurotransmitter receptors or growth factor-stimulated receptor tyrosine kinases. In neurons, known to express TRPC4 or TRPC5 together with TRPC1, like hippocampal (Chung et al., 2006) or cortical neurons (Alzheimer, 1994; Haj-Dahmane and Andrade, 1996, 1999), muscarinic receptor stimulation results in activation of a voltage-dependent nonselective cation current, which displays some similarities to currents seen following heterologous coexpression of TRPC1 with TRPC4 or TRPC5 (Strubing et al., 2001). Moreover, it has been reported that excitatory postsynaptic currents (EPSCs) in cerebellar purkinje cells, evoked by activation of metabotropic glutamate receptors (mGluR1), are mediated by TRPC1 channels (Kim et al., 2003). Together, these and other findings suggest an important role for TRPC channels in synaptic communication via endogenous neurotransmitters (Gee et al., 2003; Tozzi et al., 2003; Bengtson et al., 2004).

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[Ca]i plays an essential role in guidance of the highly organized palm-like growth cone of developing neurons by regulating growth cone morphology (Gomez and Spitzer, 1999; Song and Poo, 1999; Gomez and Spitzer, 2000; Gomez et al., 2001), the cytoskeleton (Dent and Gertler, 2003; Henley and Poo, 2004), and trafficking of membrane precursor vesicles (Craig et al., 1995). Neurite extension and growth cone steering in a gradient of netrin-1 can be inhibited by blocking plasma membrane Ca2+ channels (Mattson and Kater, 1987; Hong et al., 2000), whereas activation of voltage-dependent Ca2+ channels (Cav) in the growth cone promotes attractive steering (Lipscombe et al., 1988; Silver et al., 1990; Davenport and Kater, 1992; Spitzer et al., 2000; Ming et al., 2001; Nishiyama et al., 2003). Recent studies showing that TRP channels generate [Ca2+]i signals in growth cones strongly suggest that TRP channels are involved in Ca2+ signaling in these neuronal structures. TRPC5 channels have been shown to be enriched in the growth cones of cultured mouse hippocampal neurons where they are involved in mechanisms controlling neurite extension and growth cone morphology (Greka et al., 2003). Overexpression of TRPC5 or a dominantnegative pore mutant of TRPC5 inhibits or enhances the neurite outgrowth of the transfected neurons, respectively. Ca2+ influx through TRPC channels may also contribute to BDNF-induced elevation of [Ca2+]i and cone attraction in cultured rat cerebellar granule cells (Li et al., 2005). The BDNF receptor TrkB activates PLCg, which induces TRPC-mediated Ca2+ influx and growth cone attraction toward BDNF. TRPC1, TRPC3, and TRPC6 are expressed in granular cells, and knockdown of endogenous TRPC3 using short interfering RNA or by overexpression of dominant negative forms of TRPC3 or TRPC6 abolished BDNF growth cone attraction in the cerebellar neurons (Li et al., 2005). Interestingly, TRPC3 knockdown did not impair neurite outgrowth, indicating that Ca2+ influx through TRPC3 selectively controls growth cone guidance. Therefore, with respect to TRPC5 affecting neurite extension, different TRP channels may subserve distinct regulatory functions in the growth cone by generating different spatiotemporal patterns of Ca2+ influx. Because TRPC channels seem to be crucially involved in growth cone guidance and in the signaling via metabotropic glutamate receptors, TRPC channels, in principle, could be important in the development of many neurological diseases. Recent studies indicate a potential link between TRPC1 and neurotoxicity induced by the exogenous agent 1-methy-4-phenylpyridium (MPP+) (Bollimuntha et al., 2005). The latter causes selective nigral dopaminergic lesions and induces Parkinson disease-like syndromes. When applied to human dopaminergic SH-SY5Y neuroblastoma cells, MPP+ causes decreased expression and plasma membrane localization of TRPC1. In an opposite manner, TRPC1 overexpression reduces the neurotoxicity of MPP+, inhibits cytochrome C release by MPP+, and decreases Bax and Apaf-1 protein levels, indicating an inhibition of degenerative apoptosis involved in Parkinson’s disease. Thus TRPC1 may execute a neuroprotective role in dopaminergic neurons. Some indications suggest a role for TRPC6 in Alzheimer’s disease. Mutations in the presenilin (PS) genes are linked to the development of early-onset Alzheimer’s and transient expression of PS mutants (N141I, M239V) along with TRPC6 in human embryonic kidney (HEK)-293 cells results in a strong inhibition of agonist-induced Ca2+ entry (Lessard et al., 2005). This effect does not result from an increased secretion of amyloid b-peptides into the medium, and TRPC6 itself remains functional as evidenced by unchanged direct activation by OAG. Interestingly, the loss-of-function PS mutant D263A augments the activity of TRPC6.

2.2 TRPV The TRPV subfamily consists of six mammalian members (TRPV1, TRPV2, TRPV3, TRPV4, TRPV5, and TRPV6) (Pedersen et al., 2005; Owsianik et al., 2006). With regard to Ca2+ conductance, TRPV channels can be divided into two groups: (1) TRPV1–TRPV4 with moderate Ca2+ permeability (PCa2+/PNa+100) (Vennekens et al., 2000; Hoenderop et al., 2001; Yue et al., 2001; Voets et al., 2004b). TRPV channels are expressed in excitable and nonexcitable cells in a wide range of tissues where they have crucial roles in physiological functions and in several diseases (for review, see Levine and Alessandri-Haber, 2007; Nilius, 2007). The vanilloid receptor 1, TRPV1 (also known as VR1

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or OTRPC1), is activated by several stimuli, including heat (>43
C), capsaicin, various vanilloid compounds, camphor, allicin, NO, protons, arachidonic acid metabolites, lipoxygenase products, endocannabinoids, and 2-APB (Caterina et al., 1997; Watanabe et al., 2002b; Iida et al., 2003; Watanabe et al., 2003; Hu et al., 2004; Vriens et al., 2004b) (for review, see Tominaga and Tominaga, 2005). TRPV1 is mainly expressed in nociceptors, a subpopulation of small diameter primary afferent neurons of the dorsal root and trigeminal nerves of the PNS (Caterina et al., 1997; Tominaga et al., 1998). This ion channel is implicated in pain sensation and hypersensitivity to noxious heat (Caterina et al., 2000; Davis et al., 2000). Activation of TRPV1 in peripheral nerve endings by agonists like capsaicin promotes influx of Na+ and Ca2+, thereby, depolarizing the cells resulting in the release of neuropeptides like substance P and calcitonin gene-related peptide (CGRP) (Wood et al., 1988; Zeilhofer et al., 1997). Subsequent formation of the Ca2+–calmodulin (Ca/Cam) complex leads to receptor desensitization, suggesting a negative feedback on the channel gating. This mechanism of inactivation may underlie analgesic effects of capsaicin that are exploited for medical treatment of pain (Rosenbaum et al., 2004). A recent report showed that olvanil, a nonpungent TRPV1 agonist, downregulates voltage-activated Ca2+ channels (VACCs) through specific activation of TRPV1 in dorsal root ganglion (DRG) neurons, thereby influencing neuropeptide release (Wu et al., 2006). The effect of olvanil is mediated by activation of TRPV1 channels, which results in increased Ca2+ influx and Ca2+ release from intracellular stores. The increased [Ca2+]i initiates a complex cascade of intracellular signaling events, of which Ca2+-dependent calmodulin and calcineurin are important in the downregulation of VACCs. In central terminals of nociceptive primary afferents in the rat dorsal horn, activation of TRPV1 increases glutamate release, although this does not occur through influx of Ca2+ via activated TRPV1 channels, as capsaicin was still able to increase the miniature excitatory postsynaptic currents (mEPSCs) frequency in the absence of extracellular Ca2+ (Yang et al., 1998; Nakatsuka et al., 2002; Baccei et al., 2003). Besides its role in sensory neurons where TRPV1 mRNA is present in high concentrations (Sanchez et al., 2001), TRPV1 seems to be involved in several brain functions. TRPV1 is abundantly expressed in cell bodies and dendrites of neurons of the hippocampus and cortex and was also detected in the cerebellum, the olfactory bulb, the mesencephalon, and the hindbrain as well as in nonneuronal cell types in the brain (astrocytes and pericytes) (Toth et al., 2005; Cristino et al., 2006). TRPV1 activation in the CA1 region of the rat hippocampus enhances paired-pulse depression. It has been suggested that the mechanism underlying this depression is related to the activation of TRPV1 receptors, which cause a selective increase of inhibitory GABAergic synaptic transmission in the hippocampus (Al-Hayani et al., 2001; Huang et al., 2002). By contrast, TRPV1 activation in synaptosomal hippocampus preparations inhibited K+-evoked Ca2+ entry and reduced Ca2+-dependent K+-evoked GABA release (Kofalvi et al., 2006). The discrepancy between the in vitro and ex vivo data might be attributed to the disruption of intracellular or extracellular molecules under ex vivo conditions. Further investigation is needed to elucidate the importance of TRPV1 activation in the hippocampus and its possible role in synaptic transmission. However, a recent study implies a crucial role for TRPV1 in long-term potentiation of excitatory postsynaptic potentials (EPSPs) in the hippocampus and behavioral assays revealed a role in the regulation of anxiety and conditioned fear (Marsch et al., 2007). Evidence exists that TRPV1 is also expressed in the organum vasculosum lamina terminalis (OVLT), an area in the brain that is important for the regulation of fluid and electrolyte balance. Hypertonic stimulation of OVLT neurons leads to increased inward currents, depolarization of osmoreceptor potentials and increased action potential discharge. This signal transduction cascade is lacking in TRPV1/ mice, suggesting that TRPV1 has a role in osmosensory transduction (Ciura and Bourque, 2006). The midbrain periaqueductal gray (PAG) area is an important part of the brain for physiological functions, such as behavioral reactions, cardiovascular regulation, and pain modulation. TRPV1 activation in the dorsolateral PAG leads to increased intraterminal Ca2+ concentrations through Ca2+ influx into the nerve terminal, resulting in glutamate increase, which might have a role in the modulation of cardiovascular responses (Xing and Li, 2007). In hypothalamic slices, application of capsaicin evokes glutamate release (Sasamura et al., 1998; Sasamura and Kuraishi, 1999). The application of capsaicin in the somatosensory cortex and anterior cingulated cortex (ACC) neurons, which are involved in processing pain, slightly

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increases the membrane potential and induces an increase in action potentials. This result indicates functional expression of TRPV1 receptors in the cortical brain region (Sasamura and Kuraishi, 1999). In addition to its role as an integrator of noxious stimuli in the periphery, TRPV1 is probably involved in neurodegenerative mechanisms occurring during stroke (changes in brain temperature, pH, modulation of glutamate release) (Marinelli et al., 2002). In this context, TRPV1 agonists that promote fast desensitization and TRPV1 antagonists have been proposed to exhibit neuroprotective properties. In a model of global cerebral ischemia in gerbils, capsaicin reduced ischemia-induced hyperlocomotion, memory impairment, and electroencephalogram changes and improved survival of CA1 pyramidal cells in the hippocampus measured at various time points after the insult. Such protective effects were reduced by capsazepine (Pegorini et al., 2005). In a subsequent study employing the same model, the CB1 receptor antagonist rimonabant was also found to exert neuroprotective effects that, on the basis of antagonism by capsazepine, were at least partially attributed to the involvement of TRPV1 (Pegorini et al., 2006). One proposed mechanism is that block of CB1 by rimonabant promotes TRPV1 activation by endogenously generated compounds, such as anandamide, which in turn cause TRPV1 desensitization (Pegorini et al., 2006). However, much of the argument for the involvement of TRPV1 hinges on suppression of neuroprotection by capsazepine, which, it must be emphasized, is an agent with limited selectivity for TRPV1. Nonetheless, capsazepine has also been shown to exert a neuroprotective action in ouabain-induced neurodegeneration, but in that study CB1 receptor agonism (mediated by anadamide), rather than antagonism, was found to produce a beneficial outcome (Veldhuis et al., 2003). Schizophrenia is accompanied by morphological changes in the brain that include enlarged ventricles, a reduced cortical thickness, but an increased prefrontal neuron density. Patients also exhibit reduced pain sensitivity and a diminished niacin skin flare response. The latter symptoms hint toward a defect in TRPV1expressing afferent nerve fibers. Capsaicin treatment of neonatal rats, surprisingly, causes brain changes that resemble those found in schizophrenic patients, indicating a possible role of TRPV1 during development in the pathogenesis of this disease (Newson et al., 2005). TRPV2 is expressed in DRG neurons (Caterina et al., 1999; Ma, 2001), trigeminal ganglion (TG) neurons (Ichikawa and Sugimoto, 2000), and in various regions of the brain, including the hypothalamic paraventricular, the suprachiastic, and the supraoptic nuclei (Lewinter et al., 2004; Wainwright et al., 2004). The channel is activated by several stimuli, such as hypo-osmolarity, 2-APB, and heat (>52
C). A recent study showed that TRPV1 and TRPV2 are colocalized in vivo in DRG neurons and the cerebral cortex and form heteromultimeric channels in vitro (Liapi and Wood, 2005). TRPV3 has been identified as a heat-activated and Ca2+-permeable channel. It is expressed in superior cervical ganglia (SCG), DRG, and TG neurons, and in several brain regions (Peier et al., 2002b; Smith et al., 2002; Xu et al., 2002). TRPV3 is activated at physiological temperatures (32–39
C). Furthermore, this channel can be stimulated by camphor, carvacrol, eugenol, thymol, 2-APB, nitric oxide, arachidonic acid, and unsaturated fatty acids (Peier et al., 2002b; Smith et al., 2002; Xu et al., 2002; Chung et al., 2004; Hu et al., 2004; Moqrich et al., 2005; Hu et al., 2006; Xu et al., 2006; Yoshida et al., 2006; Vogt-Eisele et al., 2007). TRPV3 has been implicated in hyperalgesia in inflamed tissues (Hu et al., 2006; Xu et al., 2006). In dopaminergic neurons of the substantia nigra pars compacta, a region that is important in motor and emotional control, activation of TRPV3 increases the baseline [Ca2+]i. This suggests that TRPV3 is involved in the homeostasis of Ca2+ in these neurons (Guatteo et al., 2005). TRPV4 is a nonselective cation channel (Liedtke et al., 2000; Strotmann et al., 2000; Wissenbach et al., 2000; Delany et al., 2001) that is expressed in a broad range of tissues, as well as in sensory ganglia and several brain regions (Liedtke et al., 2000; Strotmann et al., 2000; Fernandez-Fernandez et al., 2002; Watanabe et al., 2002a; Alessandri-Haber et al., 2003; Chung et al., 2003; Guatteo et al., 2005; Shibasaki et al., 2007). TRPV4 can be activated by several stimuli, including osmotic cell swelling, heat, synthetic ligands such as 4a-phorbol 12,13-didecanoate (4a-PDD), mechanical force, fluid viscosity, endogenous ligands, and bisandrographolide A (Liedtke et al., 2000; Strotmann et al., 2000; Wissenbach et al., 2000; Guler et al., 2002; Watanabe et al., 2002a; Gao et al., 2003; Liedtke and Friedman, 2003; Suzuki et al., 2003; Watanabe et al., 2003; Arniges et al., 2004; Andrade et al., 2005; Smith et al., 2006). TRPV4 is involved in inflammatory, thermal and mechanical hyperalgesia (Todaka et al., 2004; Alessandri-Haber et al., 2005;

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Grant et al., 2007). Activation of TRPV4 leads to cation influx and, in dopaminergic neurons, increases spontaneous firing, suggesting that TRPV4 affects the excitability of these neurons (Guatteo et al., 2005). Similar effects were observed in hippocampal neurons, where activation of TRPV4 depolarizes the membrane potential and thereby controls the neural activity (Shibasaki et al., 2007).

2.3 TRPA TRPA1, the only mammalian member of the TRPA subfamily, is a poorly selective Ca2+-permeable [PCa/PNa ¼ 0.84 (Story et al., 2003)] cation channel. TRPA1 mRNA expression was identified in a subpopulation of small diameter nociceptors of the dorsal root, trigeminal, and nodose ganglia (Story et al., 2003; Jordt et al., 2004; Kobayashi et al., 2005; Nagata et al., 2005) and has attracted attention for its potential role in nociception and inflammatory pain (hyperalgesia). TRPA1 may also serve as a detector of a variety of other sensory processes, including detection of noxious cold and mechanosensation (Story et al., 2003; Corey et al., 2004; Jordt et al., 2004; Bautista et al., 2005; Bautista et al., 2006; Kwan et al., 2006), although these roles have not yet been fully established. The activation mechanisms of TRPA1 are diverse: Pungent plant compounds like allyl isothiocyanate (AITC) from mustard or diallyl disulfide from garlic are thiol reactive substances that induce TRPA1 channel opening (Bandell et al., 2004; Jordt et al., 2004; Bautista et al., 2005; Macpherson et al., 2005) by covalent modification of cysteine residues of the channel protein (Hinman et al., 2006). In addition, the sensitivity of TRPA1 channels for pungent chemicals seems to depend on cytosolic factors like polyphospates (Kim and Cavanaugh, 2007). Following activation, the receptor remains in a desensitized, nonactivable state. In contrast, terpene alcohols like menthol or thymol, recently found to promote TRPA1 channel opening (Karashima et al., 2007), are unlikely to covalently modify proteins and are capable to repeatedly activate TRPA1. Interestingly, menthol and structurally related compounds were shown to have not only activatory but also inhibitory effects on TRPA1 channels (Macpherson et al., 2006; Karashima et al., 2007). In whole-cell and single-channel recordings of heterologously expressed TRPA1, submicromolar to low micromolar concentrations of menthol caused channel activation whereas higher concentrations lead to a reversible channel block (Karashima et al., 2007). Further, TRPA1 appears to be regulated by PLC downstream to G-protein-coupled receptors, suggesting that channel opening can be mediated by second messengers (Bandell et al., 2004; Bautista et al., 2006; Kwan et al., 2006). Bradykinin, a potent proalgesic agent associated with tissue injury and inflammation, promotes hydrolysis of membrane phosphatidylinositol bisphosphate (PIP2) by PLC activation and opens the TRPA1 channel probably by Ca2+ release from intracellular stores and Ca2+ influx from the extracellular space (Bautista et al., 2006). Whatever stimuli activates TRPA1, its response to agonists such as AITC substantially increases [Ca2+]i (Jordt et al., 2004; Nagata et al., 2005) and thereby might promote neuronal excitation or modulation of regulatory proteins. Vice versa, Ca2+ was found to exert pronounced effects on the gating of TRPA1 (Nagata et al., 2005; Doerner et al., 2007; Zurborg et al., 2007). HEK-293 cells transiently expressing TRPA1 revealed a multiphasic inward current when stimulated with AITC in the presence of a physiological external solution with Ca2+. The activation starts slowly, and then the amplitude potentiates before inactivation occurs, after which the channel remains inactivated and nonactivable. Interestingly, in the absence of extracellular Ca2+, the slowly developing inward currents do neither augment nor inactivate. When Ca2+ is suddenly added extracellularly during AITC-mediated activation, the currents augment and then inactivate, suggesting dependence of the multiphasic current characteristic on extracellular Ca2+ that has to enter the cell to exert its effects on TRPA1 channels. Indeed, two recent studies using whole-cell and excised-patch recordings suggest that intracellular Ca2+ directly gates TRPA1 channels in a PLC- and calmodulinindependent fashion, very likely via an EF-hand motive as putative Ca2+-binding site (Doerner et al., 2007; Zurborg et al., 2007). These studies also revealed that Ca2+ modulates the efficacy of some TRPA1 agonists: AITC, cinnamaldehyde, and carvacrol responses are potentiated by Ca2+ suggesting a common mechanism for these agonists, whereas Ca2+ is required as coagonist for activation by icilin by interacting directly with TRPA1 (Doerner et al., 2007). Moreover, TRPA1 activation by intracellular Ca2+ is closely linked to voltage sensitivity (Nagata et al., 2005; Zurborg et al., 2007), an interaction that has also been

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reported for the Ca2+ activated TRP channels TRPM4 and TRPM5 (Hofmann et al., 2003; Nilius et al., 2003). Release of Ca2+ from intracellular stores was also shown to activate TRPA1 (Jordt et al., 2004; Doerner et al., 2007; Zurborg et al., 2007), a finding that is still disputed as another study failed to reproduce activation of TRPA1 by store depletion (Bandell et al., 2004). Physiological stimuli that lead to an increase in intracellular Ca2+ might stimulate TRPA1 or alternatively sensitize the channel and amplify agonist provoked responses. Since TRPA1 is permeant for Ca2+, channel activation by Ca2+ represents a self-amplifying positive feedback loop that boosts the initial Ca2+ and voltage signal. The excessive Ca2+ influx is supposed to be shut down by a Ca2+-dependent desensitization process (Nagata et al., 2005) that minimizes the Ca2+ conductivity of TRPA1 in a selflimiting process that might be indispensable in order to prevent cellular Ca2+ overload. TRPA1 was not only localized in the sensory ganglia and in the peripheral nerve endings (Bautista et al., 2005; Nagata et al., 2005) which potentially come into contact with environmental stimuli, but also in central processes of sensory neurons in superficial laminae (marginal layer and substantia gelatinosa, SG) in the spinal dorsal horn. Central neurons in the SG receive nociceptive information from peripheral tissues like the skin, the viscera, and other organs through primary afferent fibers (Basbaum and Jessell, 2000). Synaptic efficacy critically depends on the presynaptic intracellular calcium concentration and synaptic release is calcium sensitive (for review, see Sudhof, 2004). TRPV1 and P2X, two ion channels that are involved in the arousal of pain, have also been localized at central terminals of nociceptors at the SG (Nakatsuka et al., 2002) and were shown to modulate synaptic transmission by enhancing glutamate release by a jet unidentified mechanism, strengthening the synaptic connection between primary afferent fibers and spinal dorsal horn neurons (Yang et al., 1998; Nakatsuka and Gu, 2001). Recently, AITC and other TRPA1 agonists were demonstrated to cause a robust glutamate release onto SG neurons in a spinal cord slice preparation resulting in increased spontaneous EPSC frequency and amplitude (Kosugi et al., 2007). Also in the presence of La3+, a blocker of voltage-gated Ca2+ channels AITC could still induce an increase of spontaneous EPSCs. AITC action was suppressed in a Ca2+-free solution indicating that AITC enhances the glutamate release by direct Ca2+ entry through TRPA1 channels in the nerve terminals.

2.4 TRPM The TRPM subfamily consists of eight members (TRPM1–TRPM8). TRPM2, TRPM6, and TRPM7 are unique among known ion channels because of the presence of an intracellular enzymatically active protein domain attached to the ion channel (for review, see Kraft and Harteneck, 2005). Most TRPM channels have a low selectivity for Ca2+ over Na+ (TRPM2, TRPM6, TRPM7, and TRPM8: PCa/PNa = 0.3–7) (Sano et al., 2001; Kraft et al., 2004; Voets et al., 2004c), whereas TRPM4 and TRPM5 are impermeable for divalent cations (PCa/PNa = 0.05). Splice variants for TRPM2, TRPM3, and TRPM4 have been observed that show different ion filter characteristics, e.g., the longer TRPM3 variant, TRPM3a2, is highly Ca2+ permeable (PCa/PNa > 100), whereas the shorter version TRPM3a1 is more selective for Na+ (PCa/PNa = 0.05) (Grimm et al., 2003; Lee et al., 2003). TRPM channels mediate cation influx, thereby raising intracellular [Ca2+]i and [Na+]i, and depolarize the membrane. Increase in [Ca2+]i affects Ca2+-dependent pathways, whereas depolarization supports Ca2+ influx through voltage-gated Ca2+ channels and influences action potential propagation. Some TRPMs are found in the mammalian CNS but their physiological role remains largely enigmatic. TRPM2 is expressed in different brain tissues such as the cerebellum, cortex, medulla, and hippocampus (Nagamine et al., 1998; Sano et al., 2001; Hara et al., 2002; Kraft et al., 2004; Uemura et al., 2005). A truncated variant of TRPM2 is expressed in the striatum (i.e., caudate nucleus and putamen) (Nagamine et al., 1998; Hara et al., 2002; Uemura et al., 2005). At its C-terminus, TRPM2 expresses a nudix hydrolase domain that is implicated in channel activation (Perraud et al., 2001; Hara et al., 2002). The channel is activated by ADP-ribose (ADPR) (by its binding to the nudix box motif), nicotinamide adenine dinucleotide (NAD), oxidative stress (especially H2O2), and arachidonic acid (Sano et al., 2001; Kraft et al., 2004). In microglial cells and striatal neurons, when activated by H2O2, TRPM2 causes a robust increase of [Ca2+]i and cation currents, which suggests an important role of this channel in calcium signaling and cation influx

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in these cells (Smith et al., 2003; Kraft et al., 2004). Additionally, calcium influx through TRPM2 channels has a positive feedback effect on channel activation (McHugh et al., 2003). TRPM2 seems to be critically involved in CNS cell death and stroke. Ca2+ influx via NMDA-R induces an activating effect on TRPM2, thereby generating a positive feedback on Ca2+ entry. Generation of ROS (reactive oxygen/nitrogen species) further activates TRPM2. Subsequent activation of DNA repair enzymes, such as poly-ADPR polymerase, or poly-ADPR glycohydrolase, produce the TRPM2-activating intracellular ligand ADPR. Furthermore, the opening of the mitochondrial permeability transition pore (mPTP) followed by NAD+ release and subsequent conversion to ADPR will also activate TRPM2. These situations will create a vicious circle leading finally to cell death and stroke (for a detailed review, see MacDonald et al., 2006). TRPM4 is a Ca2+-activated voltage-dependent monovalent cation channel regulated by PIP2 (Nilius et al., 2006) (for review, see Rohacs and Nilius, 2007). An intriguing function for TRPM4 can be anticipated from experiments in which the GABA receptor antagonist bicuculline was employed to initiate spontaneous epileptic activity in neocortical slices. During paroxysmal depolarization shift discharges, TRPM4-like channels are activated and appear to play a role in maintaining subsequent sustained afterdepolarization waveforms. The latter effect depends on an increase in [Ca2+]i and can be blocked by maneuvers that inhibit TRPM4 (Schiller, 2004). TRPM4 might also contribute to vascular stroke. During stroke in hypoxic gliotic tissue a nonselective Ca2+-activated channel (NCCa-ATP) opens under conditions of [ATP]i depletion. This channel, which shares many properties with TRPM4 (i.e., single-channel conductance, concentration range of activation by [Ca2+]i, submicromolar block by ATP, voltage-dependent open probability) is found in astrocytes in the adult brain and is regulated by SUR1. Activation of NCCa-ATP causes complete membrane depolarization and cell swelling in astrocytes from injured brains (Chen and Simard, 2001; Simard and Chen, 2004). It would be interesting to determine whether TRPM4 is indeed the channel regulated by SUR1. If so, TRPM4 might provide a new and promising therapeutic approach to stroke. TRPM5 shares the same activation mechanism and ion selectivity as TRPM4. Similar to TRPM4, TRPM5 encodes a voltage-dependent Ca2+-impermeable cation channel (Hofmann et al., 2003; Prawitt et al., 2003) and is directly activated by intracellular Ca2+ [EC50 = 0.3–1 mM], but it is inhibited by higher [Ca2+]i (Liu and Liman, 2003; Prawitt et al., 2003; Ullrich et al., 2005). TRPM5 is predominantly expressed in taste receptors, nonneuronal chemosensory cells within the taste buds of the lingual epithelium, and has an important role in taste signaling (Zhang et al., 2003; Talavera et al., 2005). Following stimulation of G-protein-coupled taste receptor proteins, the increase of the intracellular Ca2+ concentration activates TRPM5 inducing a depolarization of the taste receptor cells, which in turn leads to the transmission of the excitation to gustatory afferent fibers (Huang et al., 2007; Zhang et al., 2007). TRPM7 is a Mg2+- and Ca2+-permeable cation channel. Similar to TRPM6, TRPM7 has a higher affinity for Mg2+ than for Ca2+, and its activity is regulated by Mg2+ and Mg2+-ATP (Nadler et al., 2001; Runnels et al., 2001). Although TRPM7 shows a high affinity for Mg2+ and Ca2+, it can also permeate Mn2+ and Co2+, in addition to nonphysiological or toxic metal ions such as Ba2+, Sr2+, Ni2+, and Cd2+. This feature suggests that TRPM7 acts as an ubiquitous metal ion influx pathway (Monteilh-Zoller et al., 2003). Similar to TRPM2 and TRPM6, TRPM7 contains a protein-kinase domain. This kinase domain might have a role in channel assembly or subcellular localization (Schmitz et al., 2003; Ryazanova et al., 2004; Matsushita et al., 2005). TRPM7 activity is positively regulated by PIP2 and hydrolysis of PIP2 inhibits channel activity (Runnels et al., 2002) (for review, see Rohacs and Nilius, 2007). In cortical neurons, blockade of TRPM7 prevents Ca2+ uptake, thereby inhibiting ROS production. This inhibition prevents anoxic death. These data suggest that TRPM7 has a key role in anoxic neuronal cell death (Aarts et al., 2003). Furthermore, TRPM7 has been identified in the brain microglia. Because TRPM7 is a Ca2+permeable channel, Ca2+ influx through this channel might be important in Ca2+-dependent microglia functions (Jiang et al., 2003). TRPM7 has also been found in the membrane of synaptic vesicles of sympathetic neurons, where its activity affects acetylcholine release (Krapivinsky et al., 2006). TRPM7 also plays a universal role in Mg2+ homeostasis associated with basic cellular metabolism and activities such as cell viability and proliferation. Cases of anoxic neuronal death have been described that include the involvement of TRPM7 in cellular damage due to an imbalance of the normal physiological

Signal molecules and calcium

26

processes (Aarts et al., 2003). In situations of brain ischemia, oxygen–glucose deprivation (OGD) and excitotoxicity mediate neuronal death. The key processes involve high Ca2+ influx as a consequence of the excitotoxicity. Subsequent production of ROS consecutively activates another Ca2+ conductance, named IOGD, which is mediated by TRPM7, and results in cellular Ca2+ overload and cell death. In models of ischemic stroke (OGD, NaCN chemical anoxia), the activation of NMDA receptors (NMDA-R) provides a route for toxic Ca2+ influx, and TRPM7 probably provides an additional pathway (MacDonald et al., 2006). TRPM7 is activated by products of neuronal NO synthase (free radicals, ROS) and transient depletion of extracellular Ca2+ and Mg2+ (e.g., by a decrease in [Mg2+]i). In addition, depolarization by TRPM7 activation will relieve voltage-dependent block of NMDA-R by Mg2+, inducing a positive feedback on NMDA-R-mediated Ca2+ entry. TRPM8 has been extensively studied for its potential role in the mammalian PNS (McKemy et al., 2002; Peier et al., 2002a; Reid et al., 2002; Nealen et al., 2003; Thut et al., 2003; Abe et al., 2005). Besides its expression in a subset of DRG and TG neurons, TRPM8 is present in taste papillae and the prostate (Tsavaler et al., 2001; Abe et al., 2005). TRPM8 is a Ca2+-permeable cation channel that has low selectivity for Ca2+ over Na2+ (PCa/PNa = 0.97–3.2) (McKemy et al., 2002; Peier et al., 2002a) and can be activated by several stimuli, such as gentle cooling ( Figure 27-1). The EF‐hand motif or helix‐ loop‐helix motif was first described over 30 years ago in carp parvalbumin (Kretsinger and Nockolds, 1973). This motif can be easily visualized by the thumb and the index of a hand, the bent major finger forming the loop that provides the normally six oxygen atoms coordinating calcium (> Figure 27-1). EF‐hand motifs are found in all EF‐hand CaBP. However, not all EF‐hands are functional, i.e., they do not all bind calcium. The members of the EF‐hand superfamily can be divided into two main categories, according to their calcium affinity and their ability to change conformation following binding of calcium (da Silva and Reinach, 1991; Ames et al., 1996; Nelson and Chazin, 1998). The first group of proteins is formed of so‐ called calcium sensors that are involved in the transmission of calcium signaling: this group consists of several subgroups such as calmodulin (CaM), proteins of the S100 superfamily, or neuronal calcium sensors (NCS). The second group of proteins is constituted by the calcium‐buffering proteins including parvalbumin, calbindin 3 (previously named calbindin D9k, CaBP9k, or CABP1) (Marenholz et al., 2004), or calretinin (Heizmann and Braun, 1992; Heizmann and Braun, 1995). In this review, we will focus only on the mammalian EF‐hand calcium binding proteins involved in signal transduction in the CNS.

2

Calmodulin

CaM is the representative example of the sensor class of EF‐hand proteins (Cohen and Klee, 1988). CaM is a small protein of 148 residues ubiquitously expressed and highly conserved among species (7/150 amino acid differences between human and yeast). CaM possesses four EF‐hand calcium‐binding sites distributed in pairs separated by a long central helix (> Table 27-1). CaM binds calcium in the micromolar range (Cox et al., 1981). CaM can regulate its target proteins in a calcium‐dependent or ‐independent way. These targets have been classified into ‘‘classic’’ and ‘‘nonclassic’’ targets. The ‘‘classic’’ targets have been reviewed extensively in several books (Cohen and Klee, 1988; Van Eldik and Watterson, 1998;

Calcium regulation by EF‐hand protein in the brain

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. Figure 27-1 First EF‐hand of human recombinant CaM. The helices A and B are shown as ribbon. The residues, implicated in calcium coordination (side chains of Asp20, Asp22, and Asp24 and backbone carbonyl of Thr27) are shown as sticks. The figure was generated with PyMol using pdb file 1CLL

Carafoli and Klee, 1999). These targets include phosphorylase kinase, myosin light chain kinase, Ca2þ/Mg2þ ATPase, phosphodiesterase, Ca2þ/CaM‐dependent protein kinase II, and calcineurin (CaN). The list of the ‘‘nonclassic’’ targets of CaM has grown exponentially in the last few years because of the screening of the gene sequence database, and is well described in the review of Chun and Sacks (2000). These targets include proteins with the IQ motif first described in unconventionnal myosins (Cheney and Mooseker, 1992). Depending on the presence of other key residues than IQ in the motif, IQ‐containing proteins can bind CaM in a calcium‐dependent or ‐independent manner (Houdusse and Cohen, 1995; Houdusse and Cohen, 1996). ‘‘Nonclassic’’ CaM targets includes cell surface receptors and channels such as the insulin receptor and the cyclic nucleotide‐gated channel, intracellular signaling proteins such as the insulin receptor substrates IRS‐1 and IRS‐2 and the phosphatidylinositol‐3 kinase PI3K, small GTPase receptors such as Rab3A, and certain nuclear proteins such as p68 RNA helicase (Chun and Sacks, 2000). The existence of so many different targets for CaM indicates a key role for this protein in many cellular processes. In the rat, human, and mouse brain, three different genes have been detected which all encode an identical protein (Fischer et al., 1988; Danchin et al., 1989; Nojima, 1989). The three genes are differentially expressed in the mouse and rat brain, suggesting that precise regulation of CaM expression levels might be attained through the contribution of the three different genes (Ikeshima et al., 1993; Sola et al., 1996; Palfi et al., 2002). In the CNS, CaM plays an essential role in synaptic transmission and neuronal plasticity associated with short‐term and long‐term potentiation, and learning and memory processes, through the activation of Ca2þ/CaM‐dependent enzymes such as Ca2þ/CaM‐dependent adenyl cyclase, Ca2þ/CaM‐dependent protein kinase II, and Ca2þ/CaM‐dependent protein phosphatase 2B and through

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. Table 27-1 Characteristics of EF‐hand containing proteins (From Swiss‐Prot and TrEMBLhuman protein databases except when indicated otherwise) Protein CaM CaBP1 CaBP2 CaBP3 CaBP4 CaBP5 CaBP7 CaBP8 Recoverin NCS‐1 VILIP‐1 VILIP‐2 (rat) VILIP‐3 GCAP1 GCAP2 GCAP3 KChIP1 KChIP2 KChIP3 KChIP4 Sorcin ALG‐2 Peflin Calpain (small) Calpain (large) CaNB CHP1 CHP2 Tescalcin Calmyrin S100 B S100A1 S100A4 S100A5 S100A6 S100A10 S100A12 S100A13 S100A16 Aralar1 Citrin SCaMC a

Number of EF‐hands 4 4 (EF2 nonfunctional) 4 (EF2 nonfunctional) 3 (EF1 nonfunctional) 4 (EF2 nonfunctional) 4 (EF2 nonfunctional) 2 2 4 (EF1, ‐4 nonfunctional) 4 (EF1 nonfunctional) 4 (EF1 nonfunctional) 4 (EF1 nonfunctional) 4 (EF1 nonfunctional) 4 (EF1 nonfunctional) 4 (EF1 nonfunctional) 4 (EF1 nonfunctional) 4 (EF1 nonfunctional) 4 (EF1 nonfunctional)a 4 (EF1 nonfunctional)a 4 (EF1 nonfunctional)c 5 (EF1, ‐4,‐5 nonfunctional) 5 (EF4,‐5 nonfunctional) 5 (EF5 nonfunctional) 5 (EF5 nonfunctional) 5 (EF5 nonfunctional) 4 4 (EF1, ‐2 nonfunctional) 4 (EF1, ‐2 nonfunctional) 1 2 2 2 2 2 2 2 (nonfunctional) 2 2 2 (EF1 nonfunctional)d 4 (EF3 nonfunctional) 4 (EF3 nonfunctional) 4e

Burgoyne and Weiss, 2001 An et al., 2000 c Burgoyne, 2004 d Sturchler et al., 2006 e del Arco and Satrustegui, 2004 b

MW (kDa) 17 25.8 24.3 21.7 30.4 19.8 24.4 24.8 23 21.7 22 22.1 22.2 22.7 23.3 23.6 23 27–28b 27–28b 26.5 21.6 21.8 30.3 19 80 19.4 22.3 22.3 24.7 22 10.5 10.4 11.7 10.7 10.1 11 10.4 11.4 11.8 74.7 74.1 52.3

Myristoylation site No Yes Yes No No No No No Yes Yes Yes Yes Yes Yes Yes Yes Yes No (N‐ter hydrophobic sequence) No (N‐ter hydrophobic sequence) No (N‐ter hydrophobic sequence) No (N‐ter hydrophobic sequence) No (N‐ter hydrophobic sequence) No (N‐ter hydrophobic sequence) No (N‐ter hydrophobic sequence) No (N‐ter hydrophobic sequence) Yes Yes Yes No Yes No No No No No No No No No No No No

Calcium regulation by EF‐hand protein in the brain

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the interaction with other regulatory proteins such as Munc13 (Onyike et al., 1998; Zeng et al., 2001; Colbran, 2004; Junge et al., 2004; Matsuzaki et al., 2004). Calcium‐loaded CaM interacts directly with RC3 or neurogranin, a protein suggested to play an important role in postsynaptic neuronal functions (van Dalen et al., 2003). Ca2þ/CaM and Ca2þ/CaM‐dependent protein kinase may also be involved in memory and motor deficits associated with lead exposure (Gill et al., 2003). Ca2þ/CaM may be involved in Parkinson’s disease by interacting with a‐synuclein (> Table 27-2) and in neurodegenerative diseases in general by regulating the death‐associated protein kinase (DAPK) (Martinez et al., 2003; Schumacher et al., 2004). . Table 27-2 Association of EF‐hand proteins with some brain pathologies Protein Calmodulin S100B

S100A6 NCS‐1 VILIP‐1 KChIp4 Calcineurin

Sorcin Calpain

3

Disease Parkinson’s disease Down’s syndrome Alzheimer’s disease Epilepsy Depression Schizophrenia Amyotrophic Lateral Sclerosis Schizophrenia Bipolar disorder Alzheimer’s disease Alzheimer’s disease Amyotrophic Lateral Sclerosis Alzheimer’s disease Alzheimer’s disease Alzheimer’s disease

Reference Martinez et al. (2003) Allore et al. (1988) Sheng et al. (1994) Dyck et al. (2002) Schroeter et al. (2002) Lara et al. (2001) Hoyaux et al. (2002) Koh et al. (2003) Schnurra et al. (2001) Morohashi et al. (2002) Ferri et al. (2004) Ermak et al. (2001); Hata et al. (2001); Lian et al. (2001) Pack‐Chung et al., 2000) Veeranna et al., 2004

The S100 Protein Superfamily

The S100 protein superfamily is the largest subgroup of EF‐Hand‐containing proteins and is now constituted of 21 members (Heizmann et al., 2002; Marenholz et al., 2004). S100 proteins are small (10–12 kDa) acidic proteins and contain two EF‐hands (> Table 27-1). A canonical EF‐hand is located at the C‐terminus whereas the N‐terminus contains an EF‐hand specific for each S100 protein (Bhattacharya et al., 2004; Fritz and Heizmann, 2004). The affinity for calcium ranges from 20–500mM (Heizmann and Cox, 1998). In solution, the proteins form principally homo‐ or heterodimers, except for the single member calbindin 3, occurring only as monomer (Fritz and Heizmann, 2004). In addition, calbindin 3 is the only member of the S100 family to act as a calcium buffer and not a sensor. S100 proteins are restricted to vertebrates and are almost all (16/20) located in region 1q21 of the human chromosome 1 (Marenholz et al., 2004). Interestingly, the gene coding S100B is located on human chromosome 21, linking this family member to brain diseases such as Alzheimer’s disease (AD) (Van Eldik and Wainwright, 2003) or Down’s syndrome (> Table 27-2) (Allore et al., 1988). Among the 21 known members of this family, not all of them are present in the CNS. > Table 27-3 summarizes the characteristics of the S100 proteins expressed in the CNS.

3.1 S100A1 S100A1 and S100B were the first two proteins described in the family and are best characterized (Moore, 1965). Although S100A1 is mainly expressed in skeletal and cardiac muscle, it is also found in the brain (Zimmer, 1991). S100A1 interacts in vitro with more than 10 various proteins or enzymes and has

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. Table 27-3 Characteristics of S100 protein members expressed in the CNS (for nomenclature see Marenholz et al., 2006) Protein S100B

Distribution in the CNS Entorinal, occipital cortex, hippocampus (Tiu et al., 2000) Neurons, astrocytes, oligodendrocytes (Van Eldik and Wainwright, 2003)

S100A1

Neurons (Benfenati et al., 2004)

S100A4

Sensory and autonomic neurons, Schwann cells (Sandelin et al., 2004), White matter astrocytes (Kozlova and Lukanidin, 1999; Aberg and Kozlova, 2000), Optic nerve (Aberg and Kozlova, 2000), Developing human hippocampus and temporal cortex (Chan et al., 2003)

Target proteins Neuromodulin GAP‐43 (Lin et al., 1994) Tau (Baudier and Cole, 1988) GFAP (Ziegler et al., 1998; Frizzo et al., 2004) Vimentin, microtubules, intermediates filaments type III (Sorci et al., 1998) Annexin VI (Garbuglia et al., 1998; Garbuglia et al., 2000; Arcuri et al., 2002) TRTK‐12 (Frizzo et al., 2004; McClintock et al., 2002; Inman et al., 2002) p53 (Delphin et al., 1999) sgt‐1 (Nowotny et al., 2003) CacyBP/BP (Filipek et al., 2002) Ndr protein kinase (Millward et al., 1998) AHNAK (Gentil et al., 2001) Phosphoglucomutase (Landar et al., 1996) Fructose 1,6‐biphosphatase, aldolase (Zimmer and Van Eldik, 1986) RAGE (Hofmann et al., 1999) Calponin (Fujii et al., 1994) Caldesmon (Skripnikova and Gusev, 1989) Neurocalcin (Okazaki et al., 1995) CapZ (Ivanenkov et al., 1995) Fructose 1,6‐biphosphate, aldolase (Zimmer and Van Eldik, 1986) Glycogen phosphorylase (Zimmer and Dubuisson, 1993) Adenylate cyclase (Fano et al., 1989); Phosphoglucomutase (Landar et al., 1996) Tubulin (Donato, 1988) GCAP (Ivanenkov et al., 1995) MyoD (Baudier et al., 1995) Intermediate filaments type III (Garbuglia et al., 1999) annexin VI (Garbuglia et al., 1998) synapsin I (Heierhorst et al., 1999; Benfenati et al., 2004) Ryanodine receptor (Treves et al., 1997) Twitchin kinase (Heierhorst et al., 1997) SERCA‐2a (Kiewitz et al., 2003; Most et al., 2005) RAGE (Huttunen et al., 2000) Hsp70, Hsp90, FKBP52, CyP40 (Okada et al., 2004) Titin (Yamasaki et al., 2001) Nonmuscle tropomyosin (Takenaga et al., 1994); p53 (Parker et al., 1994); non‐muscle myosin (Ford and Zain, 1995; Kim and Helfman, 2003); Map (Watanabe et al., 1992)

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. Table 27-3 (continued) Protein S100A5

S100A6

Distribution in the CNS Olfactory bulb, brainstem, spinal trigeminal tract (Schafer et al., 2000) Astrocytic tumors (Camby et al., 1999) Developing human hippocampus and temporal cortex (Chan et al., 2003) Subpopulation of neurons, astrocytes, (Yamashita et al., 1999) Entorinal, occipital cortex, hippocampus (Tiu et al., 2000)

S100A10

Brain (Allen et al., 1996)

S100A13

Amygdala, striatum, cerebellum, cerebral cortex Hippocampus, substantia nigra, thalamus, spinal cord (Ridinger et al., 2000) Developing hippocampus and temporal cortex (Chan et al., 2003) Brain (Marenholz and Heizmann 2004) Cerebral cortex, basal ganglia, hippocampus Amygdala, thalamus, hypothalamus, cerebellum (Sturchler et al., unpublished)

S100A16

Target proteins No target described yet

Glyceraldehyde 3‐phosphate dehydrogenase (Filipek et al., 1995), CacyBP/SIP (Nowotny et al., 2003; Filipek et al., 2002) Annexin I, II, VI, XI, (Sudo and Hidaka 1998; Filipek et al., 1995; Watanabe et al., 1993; Zeng et al., 1993) p30 (Filipek and Wojda 1996) Annexin II (Benaud et al., 2004; van de Graaf et al., 2003; Zobiack et al., 2003) Phospholipase A2 (Wu et al., 1997) Tissue‐type plasminogen activator (Kang et al., 1999) Tetrodotoxin‐resistant sodium channel (Okuse et al., 2002) Synaptotagmin 1 (Landriscina et al., 2001)

No target described yet

been described to be involved in regulation of energy metabolism, protein phosphorylation, regulation of cytoskeleton constituents, or muscle contraction (> Table 27-3). S100A1 is expressed during neuronal differentiation as shown in cell culture experiments (Zimmer and Landar, 1995). Although the secretion of S100A1 has not yet been demonstrated, it exerts trophic effects on neurons in vitro via a receptor suggested to be receptor for advanced glycation endproducts (RAGE) (Huttunen et al., 2000). Recent experiments have also suggested that S100A1 may play a role as a molecular chaperone in a complex with Hsp70/Hsp90 (Okada et al., 2004).

3.2 S100B S100B protein represents 0.2% of total brain protein and is mainly synthesized by astrocytes, oligodendrocytes, and Schwann cells and to a lesser extent by neurons (Tiu et al., 2000; Chan et al., 2003; Donato, 2003; Zimmer et al., 2003). S100B has been shown to interact in vitro with more than 15 proteins involved in various cellular functions such as cell metabolism, cell motility, cell death, or synaptic plasticity (see > Table 27-3). S100B is also secreted by astrocytes and exerts either a trophic or a toxic effect on neurons depending of its concentration: at nanomolar concentration, S100B stimulates neurite outgrowth and

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enhances survival of neurons during development and after glucose deprivation damage, whereas a micromolar concentration of S100B induces apoptosis (Van Eldik et al., 1991; Barger et al., 1995; Huttunen et al., 2000). As with S100A1, RAGE has been suggested to be the multiligand receptor mediating the extracellular effects observed with S100B (Hofmann et al., 1999). S100B protein is not an essential protein for survival since both S100B knockout mice and transgenic animals overexpressing S100B present only mild phenotypes. S100B knockout mice do not show any obvious abnormalities in development, although some animals show an enhanced spatial and fear memory associated with strengthened neuronal plasticity. In addition, enhanced epileptogenesis has been reported, probably because of a decrease in calcium‐ handling capacity in astrocytes (Xiong et al., 2000; Nishiyama et al., 2002). Furthermore, transgenic mice overexpressing S100B exhibit enhanced explorative activity, reduced anxiety, and impaired learning and memory capabilities (Gerlai et al., 1995; Gerlai and Roder, 1996; Winocur et al., 2001; Bell et al., 2003). Interestingly, in humans higher S100B concentrations have been detected after brain trauma and ischemia, making S100B a potential diagnostic protein in traumatic brain damage (Rothermundt et al., 2003). Increased concentration of S100B has also been found in cases of neurodegenerative disease, such as Alzheimer’s disease (Griffin et al., 1989; Mrak et al., 1996), in patients with Down syndrome, where a 1.7‐fold increase in the concentration of S100B is found (Mito and Becker, 1993; Griffin et al., 1998), and in some psychiatric disorders such as schizophrenia (> Table 27-2).

3.3 S100A4 S100A4 protein is involved in cell cycle regulation and is closely associated with metastatic activity of some malignant tumors (> Table 27-3) (Barraclough, 1998; Gupta et al., 2003). S100A4 was also found to be upregulated in astrocytes of sectioned spinal cord in adult rats, both at the mRNA and the protein level, suggesting a role of S100A4 in astrocytic responses to injury (Zhang et al., 2004). S100A4 can also induce neurite extension through activation of the ERK1/2 signaling pathway in neurons (Novitskaya et al., 2000).

3.4 S100A5 S100A5 is expressed in certain areas of the brain, namely, the olfactory bulb, the brainstem, and the spinal trigeminal tract (> Table 27-3) (Camby et al., 2000; Schafer et al., 2000). It is also overexpressed in astrocytic tumors (Camby et al., 2000) and has been suggested to be a marker of recurrence in certain meningiomas (Hancq et al., 2004). No target protein for S100A5 has been yet described.

3.5 S100A6 As with S100A5, the next member of the family, S100A6, is also restricted to some subpopulations of neurons and astrocytes (Yamashita et al., 1999). S100A6 is overexpressed in astrocytes associated with the neurodegenerative lesions of amyotrophic lateral sclerosis (ALS) (> Table 27-2) (Hoyaux et al., 2002). It was also found recently to be upregulated in astrocytes of patients with Alzheimer’s disease, and also in two different mouse models of AD (Boom et al., 2004).

3.6 S100A10 S100A10, formerly annexin II light chain or p11, is expressed in many tissues, including brain (> Table 27-3) (Allen et al., 1996). It has been shown to stimulate the conversion of plasminogen to plasmin in vitro in a calcium‐dependent manner (Kang et al., 1999; Donato, 2003; Kwon et al., 2005). It has also been found to regulate sensory neuron‐specific sodium channel expression (Okuse et al., 2002).

Calcium regulation by EF‐hand protein in the brain

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3.7 S100A12 S100A12 is mainly expressed in human granulocytes, keratinocytes, and psoriatic lesions. However, human recombinant S100A12 was found to promote neurite outgrowth in vitro (Mikkelsen et al., 2001; Moroz et al., 2003), suggesting that S100A12 plays a role in neuronal differentiation.

3.8 S100A13 S100A13 is present in many regions of the brain, including the hippocampus, cerebral cortex, cerebellum, and thalamus (> Table 27-3) (Ridinger et al., 2000). Immunoreactivity against S100A13 was also shown in the hippocampus and cerebral cortex of the human fetus at early stages of development (12–24 weeks development), suggesting a role of S100A13 in the development of human brain (Chan et al., 2003). S100A13 has been shown to be part of a multiprotein aggregate complex containing synaptotagmin 1 (Landriscina et al., 2001).

3.9 S100A16 S100A16 is the last discovered member of the S100 protein family and, interestingly, it is ubiquitously expressed (> Table 27-3) (Marenholz and Heizmann, 2004). In situ hybridization experiments performed on mouse brain sections show a widespread distribution of the S100A16 transcript in many brain areas, such as the cerebral cortex, amygdala, basal ganglia, hippocampus, thalamus, hypothalamus, and cerebellum (> Figure 27-2a); in contrast, S100B exhibit a more restricted expression pattern (> Figure 27-2b) (Sturchler et al., 2006).

. Figure 27-2 In situ hybridization analysis of S100mRNA distribution in mouse brain. Coronal cryostat sections were hybridized with a probe specific for S100A16 (a), S100B (b), or with a sense probe specific for S100A16 (c, control). HC, hippocampus; CC, corpus callosum; LGP, lateral globus pallidus; VMH, ventromedial hypothalamic neuron; Amy, amygdala. Scale bar ¼ 2.5mm

3.10 Interaction with RAGE Several members of the S100 protein superfamily are potential ligands of the RAGE (Schmidt et al., 2001). RAGE is involved in a large number of pathophysiological processes, e.g., immune/inflammatory disorders, Alzheimer’s disease, tumorigenesis, abnormalities associated with diabetes such as arteriosclerosis or

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disordered wound healing (Hori et al., 1995; Hofmann et al., 1999; Schmidt et al., 1999; Hofmann et al., 2002; Schlueter et al., 2003; Arancio et al., 2004;). The S100 proteins activating RAGE include S100A1, S100B, and S100A12 (Hofmann et al., 1999; Huttunen et al., 2000). Following their interaction with RAGE, various cellular pathways are activated involving p21Ras, endothelin 1, cdc42, MEK (ERK1/ERK2), MAP kinases, and the transcription factor NF‐kB (Schmidt et al., 1994; Yan et al., 1994; Huttunen et al., 1999; Ostendoip et al., 2007; Lecleic et al., 2007). Activation of RAGE enhances the expression of the receptor itself and initiates a positive feedback loop resulting in sustained RAGE upregulation, which has been suggested to be the starting point of chronic cellular activation and tissue damage. Recent work also suggests that RAGE is involved in axon regeneration via its interaction with S100A12/calgranulin (Rong et al., 2004).

4

The Neuronal Calcium Sensors

This family of small CaBP constitutes a fast‐growing family with more than 40 members, including 14 members in mammals (Braunewell and Gundelfinger, 1999; Burgoyne, 2004; Burgoyne et al., 2004). This family regroups several subgroups according to their amino acid similarity (Braunewell and Gundelfinger, 1999). The NCS are primarly expressed in retinal photoreceptor and neuroendocrine cells, suggesting a specialized role in these cells (Braunewell and Gundelfinger, 1999). The NCS have been involved in large variety of cellular functions, including phototransduction, regulation of neurotransmitter release, the control of cyclic nucleotide and phosphoinositide metabolism, gene expression, regulation of ion channels, and also membrane traffic and apoptosis (Mercer et al., 2000; Burgoyne and Weiss, 2001; Ivings et al., 2002; Hilfiker, 2003; Burgoyne et al., 2004). The NCS have two pairs of EF‐hand and unlike CaM and S100 proteins, possesses a consensus myristoylation sequence at the N‐terminal responsible for the targeting of the NCS to the membrane in a calcium‐dependent or ‐independent manner (> Table 27-1) (Ames et al., 1997; Valentine et al., 2003). From the four EF‐hands, the first one, EF1, is unable to bind calcium because of a mutation in the first calcium‐binding loop. All NCS members bind calcium with submicromolar affinity (reviewed in Burgoyne and Weiss, 2001).

4.1 The Recoverins The first subgroup of NCS is formed by the recoverins, including visinin in chicken, S‐modulin and S26 in frog, and recoverin expressed in various species including humans (Braunewell and Gundelfinger, 1999). The first family member described was visinin in chicken retina (Yamagata et al., 1990). Visinin is a 24 kDa protein expressed in the retinal photoreceptor layer and in pinealocytes (Goto et al., 1990; Yamagata et al., 1990). Recoverin is a 23 kDa myristoylated protein found in normal conditions only in photoreceptor cells (rods and cones) (Dizhoor et al., 1991; Ray et al., 1992). In the absence of calcium, the myristoyl group is sequestered within a hydrophobic pocket of the protein, rendering the protein cytosolic (Tanaka et al., 1995). In the presence of calcium, the protein changes its conformation, resulting in the extrusion of the myristoyl group and the insertion of the protein to the membrane, a phenomenon described as the myristoyl switch (Ames et al., 1997). The main function of recoverin is to bind and inhibit rhodopsin kinase, thereby prolongating the light response (Chen et al., 1995; Burgoyne and Weiss, 2001; Burgoyne, 2004). Also principally expressed in retinal rods, recoverin has been found to be aberrantly expressed in malignant tumors localized outside the nervous system (Bazhin et al., 2003; Bazhin et al., 2004a, b). This abnormal expression of recoverin in tumor tissue can trigger the immune system to produce autoantibodies against recoverin that might result in the degeneration of the retina caused by paraneoplastic retina degeneration or cancer‐associated retinopathy (Bazhin et al., 2004a, b).

4.2 The Guanylate Cyclase‐Activating Protein A second class of NCS is formed by the guanylate cyclase‐activating protein (GCAP) in vertebrates and the frog ortholog GCIP (Braunewell and Gundelfinger, 1999). The GCAPs are expressed only in the photoreceptor cells of the retina of vertebrates (Gorczyca et al., 1994) (reviewed by Palczewski et al., 2004).

Calcium regulation by EF‐hand protein in the brain

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They stimulate guanylate cyclase in the absence of calcium and inhibit the enzyme in the presence of calcium. There are three GCAP in vertebrates, GCAP‐1, ‐2, and ‐3. They all have four EF‐hand motifs with only three functional (> Table 27-1) (EF‐2, EF‐3, and EF‐3) (Palczewski et al., 2004).

4.3 Visinin‐like Protein In humans, the visinin-like proteins (VILIPs) regroup the VILIP‐1, VILIP‐2, hippocalcin, neurocalcin, and VILIP‐3, also named hippocalcin‐like 1 protein in the GeneBank (> Table 27-1) (Spilker et al., 2000; Spilker et al., 2002). VILIPs are found in different cell types in the cerebellum (Spilker et al., 2000). VILIP‐1 is predominantly expressed in neurons of the granule layer of the cerebellum, whereas VILIP‐3 is expressed in the Purkinje cell layer (Spilker et al., 2000). Within the same neuron, these two proteins present different calcium‐dependent translocation, suggesting higly selective response to various stimuli within the same cells (Spilker and Braunewell, 2003). VILIP‐1 was found to be associated with fibrillar tangles in Alzheimer’s brains and may have a role in tau hyperphosphorylation (> Table 27-2) (Schnurra et al., 2001).

4.4 The Frequenin/NCS‐1 A fourth group of NCS is formed by frequenin, first described in Drosophila, with the mammalian ortholog NCS‐1 (Pongs et al., 1993; Bourne et al., 2001; Strahl et al., 2003). NCS‐1 possesses four EF‐hand motifs with only three functional (EF2, 3, 4) and binds calcium with low micromolar affinity (> Table 27-1) (Cox et al., 1994). NCS‐1 has a myristoylation consensus sequence at the N‐terminus (McFerran et al., 1999). However, NCS‐1 does not follow the myristoyl switch model since the protein is able to bind to membranes in the absence of calcium (McFerran et al., 1999). NCS‐1 has been suggested to have an important role in enhancing synaptic transmission, regulating vesicular trafficking and certain voltage‐ gated ion channels (Hilfiker, 2003). Recent studies also support the hypothesis that schizophrenia and bipolar disorder may be associated with abnormalities of NCS‐1 interaction with the D2 dopamine receptor (> Table 27-2) (Koh et al., 2003). In addition, it is involved in the calcium‐dependent regulation of associative learning and memory in Caenorhabditis elegans (Gomez et al., 2001).

4.5 The Potassium Channel Interacting Proteins A fifth group of NCS is constituted by the potassium channel interacting proteins (KChIPs). These proteins modulate the activity of A‐type voltage‐gated potassium channels (An et al., 2000). Four members have been described with slightly different tissue expression pattern: whereas KchIP1 is predominantly expressed in the brain, KchIP2 is expressed in heart, brain, and lung, and KchIP3, independently discovered as DREAM or calsenilin (Carrion et al., 1999; Buxbaum, 2004), is present in brain and testis (Aberg and Kozlova, 2000; Burgoyne, 2004). A fourth member of this family has been recently characterized (Morohashi et al., 2002). The protein named KChIP4 presents 79.6% homology with the C‐terminus of KChIP2 and 77.6% homology with KChIP3, whereas its N‐terminus diverges from all other KChIP (Morohashi et al., 2002). The protein is expressed in the brain and like other members of this family, regulates the voltage‐gated potassium channel (Morohashi et al., 2002). It also interacts with PS2, suggesting a role in Alzheimer’s disease (Morohashi et al., 2002). This protein has four EF‐hands and preliminary studies indicate that it binds calcium (> Table 27-1) (Morohashi et al., 2002).

5

The CaBP

This family of EF‐hand CaBP is related to the NCS and CaM (Seidenbecher et al., 1998; Haeseleer et al., 2000). They possess 2 to 4 EF‐hand calcium‐binding sites, from which all or only some of them are functional (> Table 27-1) (Haeseleer et al., 2000). The affinity of calcium for the CaBP is not yet known.

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There are so far 7 members of CaBP described: caldendrin or CaBP1 with short and long spliced variants (Laube et al., 2002), CaBP2, 3, 4, 5, and 7 (Haeseleer et al., 2000) and the recently discovered CaBP8 (Wu et al., 2001). These proteins are expressed either only in the retina, such as CaBP2 and CaBP5, or also in the brain for the others CaBP (Haeseleer et al., 2000). The function of the CaBPs is still unclear but recent data suggest that CaBP1 regulate P/Q type calcium channels (Lee et al., 2002) and Ins(1,4,5)P3 receptors (Yang et al., 2002; Haynes et al., 2004). CaBP8 (calneuron) may play a role in the physiology of neurons and may be potentially important in memory and learning (Wu et al., 2001).

6

The Penta‐EF‐hand Family

The penta‐EF‐hand PEF family regroups a number of cytosolic proteins containing five EF‐hand domains able to translocate to the plasma membrane upon calcium binding (> Table 27-1) (Maki et al., 1997; Maki et al., 2002). In addition to the EF‐hand domains, the penta‐EF‐hand protein members have in common their propensity to dimerize via the unpaired fifth EF‐hand and to have a hydrophobic glycine/proline‐rich domain that might be involved in the translocation of the proteins to the membrane (Maki et al., 2002). To date five members of this family have been found in human tissue, sorcin, calpain light and heavy chain, grancalcin, ALG‐2, and peflin. Among these proteins, grancalcin is specifically expressed in myeloid tissue and will not be mentioned in this chapter (Teahan et al., 1992).

6.1 Sorcin Sorcin or soluble resistance‐related calcium binding protein is a 21.6 kDa protein isolated in multidrug‐ resistant cells (> Table 27-1) (Meyers and Biedler, 1981) and expressed in a few mammalian tissues such as skeletal muscle, heart, and brain. The crystal structure of sorcin showed a common folding between all the members of the PEF family (Xie et al., 2001; Ilari et al., 2002). Sorcin interacts with the ryanodine receptor in the rat caudate putamen nucleus (Pickel et al., 1997). In the rat brain, coimmunostaining experiments have shown the colocalization of sorcin with NMDA receptors in only certain NMDA‐containing neurons, suggesting differential vulnerabilities of certain neurons to excitoxin (Gracy et al., 1999). Sorcin may also be linked to Alzheimer’s disease as it interacts with part of the presenilin protein (PS2) (> Table 27-2) (Pack‐Chung et al., 2000).

6.2 Calpain Calpain belongs to the family of cytosolic calcium‐dependent cysteine proteinases (Croall and DeMartino, 1991; Jung et al., 2001; Sato and Kawashima, 2001). Calpains are involved in a large number of physiological processes such as cell cycle regulation or apoptosis, and in pathophysiological processes such as Alzheimer’s or Parkinson’s disease (Di Rosa et al., 2002; Crocker et al., 2003; Nixon, 2003; Veeranna et al., 2004). At least 14 members of the calpain family have been identified, which can be either tissue‐specific or ubiquitously expressed (Suzuki and Sorimachi, 1998). Most calpains are composed of heterodimers with a large (80kDa) catalytic subunit and a small (30 kDa) regulatory subunit. Both large and regulatory subunits contain the penta‐EF‐hand motif (> Table 27-1) (Blanchard et al., 1997; Lin et al., 1997; Carafoli and Molinari, 1998; Sorimachi and Suzuki, 2001). The two main isoforms, calpain I (mcalpain) and calpain II (mcalpain), differ in their calcium requirement: mcalpain, requires millimolar calcium concentration to be active (Mellgren and Rozanov, 1990), whereas mcalpain, which is the major form expressed in neuronal cells, is active at low micromolar calcium concentrations. Each subunit binds only four calcium ions (Michetti et al., 1997). In Alzheimer’s disease, calpain is suggested to be involved in hyperphosphorylation of neurofilaments and tau proteins that are observed at early stages of the disease (> Table 27-2) (Veeranna et al., 2004).

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6.3 ALG‐2 ALG‐2 is the first calcium‐binding protein of the EF‐hand family found to be directly involved in apoptosis (Vito et al., 1996; Tarabykina et al., 2004). ALG‐2 is a 22 kDa protein and like the other members of the penta‐EF‐hand family, contains 5 EF‐hands, with only two of them functional (> Table 27-1) (Tarabykina et al., 2000; Krebs et al., 2002). The protein binds two calcium ions in the EF‐hands 1 and 3 with low micromolar affinity (Tarabykina et al., 2000). ALG‐2 protein is expressed in mouse brain and eye and was found to be upregulated in various cancer tissues such as mammary tumors, hepatomas, and lung cancer tissue (Krebs et al., 2002; la Cour et al., 2003). Several targets have been found, such as proteins AIP, Alix, peflin, annexin VII, and annexin XI, reinforcing the putative role of ALG‐2 in apoptosis (Missotten et al., 1999; Vito et al., 1999; Kitaura et al., 2001; Satoh et al., 2002).

6.4 Peflin Peflin presents 40.9% identity with ALG‐2, particularly in EF‐1 (46.2%) and EF‐3 (57.1%) domains (> Table 27-1) (Kitaura et al., 1999). The protein has a long hydrophobic N‐terminal extension containing nine nonapeptide (A/PPGGPYGGP) repeats. Peflin has been shown to be ubiquitously expressed (Kitaura et al., 1999; Maki et al., 2002). The function of the protein peflin is still unknown although recent experiments show the formation of peflin/ALG‐2 heterodimers in certain conditions, suggesting a role in calcium‐dependent apoptosis processes (Kitaura et al., 2001).

7

Other EF‐hand CaBP in the Brain

7.1 Calcineurin Although not related to any of the family of the EF‐hand CaBP mentioned above, CaN plays essential roles in the CNS and particularly in neurons (Guerini, 1997; Groth et al., 2003). CaN or protein phosphatase 2B is a protein phosphatase expressed in many tissues but selectively enriched in neurons (Wang and Desai, 1976; Klee et al., 1979; Kincaid et al., 1987). CaN consists of a catalytic subunit (CaNA) of about 60kDa, and a regulatory subunit (CaNB) of 19kDa (Takaishi et al., 1991; Kuno et al., 1992). Activation of CaN requires both the interaction of Ca2þ/CaM to the catalytic subunit and the binding of calcium to the regulatory subunit CaNB (Stewart et al., 1982; Stemmer and Klee, 1994). CaNB possesses four functional EF‐hands which bind calcium with micromolar affinity (> Table 27-1) (Stemmer and Klee, 1994). In the brain, CaN regulates a large variety of proteins such as transcription factors, calcium channels, GABA, and glutamate receptor (reviewed in (Groth et al., 2003). CaN also regulates certain inhibitors of protein phosphatase 1 (PP1), resulting of an increase in PP1 activity, thereby regulating sodium channels and ion pumps (King et al., 1984; Gustafson et al., 1991; Ouimet et al., 1998). Besides the regulation of protein phosphorylation by PP1, CaN also directly regulates neuronal cytoskeletal proteins, enzymes involved in neurotransmitter synthesis, and neurostransmitter receptors, and also neurotransmitter release, resulting in neuronal plasticity (Groth et al., 2003). Recent data suggest that CaN may also be involved in certain neurodegenerative disorders such as ALS or Alzheimer’s disease (Ermak et al., 2001; Hata et al., 2001; Lian et al., 2001; Ferri et al., 2004).

7.2 CaN Homologous Protein CHP This family of EF‐hand‐containing proteins are homologous to CaNB. The first described member was CaN homologous protein 1 (CHP1) also known as P22 (Barroso et al., 1996). The second member, CHP2, possesses 61% identity to CHP1 (Rusnak and Mertz, 2000; Pang et al., 2002). CHP1 and CHP2 are ubiquitously distributed, and have four EF‐hand domains and one putative site of myristoylation

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(> Table 27-1). A third member of this family is tescalcin, a 24 kDa protein consisting of only one functional EF‐hand domain with three additional domains homologous to EF‐hand (Gutierrez‐Ford et al., 2003). Tescalcin binds calcium with submicromolar affinity (Gutierrez‐Ford et al., 2003). Together with CHP1 and CHP2, tescalcin possesses a site of myristoylation (Gutierrez‐Ford et al., 2003). However, the expression of tescalcin is restricted to certain tissues, including heart, brain, and stomach (Gutierrez‐ Ford et al., 2003). CHP1, ‐2, and tescalcin have been shown to inhibit the phosphatase activity of CaNA (Gutierrez‐Ford et al., 2003).

7.3 Calmyrin Calmyrin is most closely related to human CaNB, sharing 25% identity and 44% overall similarity (Stabler et al., 1999). The protein contains 2 EF‐hands and a myristoylation site at the N‐terminus (> Table 27-1) (Stabler et al., 1999). Preliminary experiments suggest that calmyrin binds calcium in vitro (Stabler et al., 1999). Calmyrin interacts preferentially with presenilin 2 (PS2) and may thus be involved in Alzheimer’s disease (> Table 27-2) (Stabler et al., 1999).

7.4 The Calcium Mitochondrial Carrier Superfamily A new subfamily of mitochondrial carriers has been discovered recently: the calcium mitochondrial carriers (CaMCs) containing EF‐hand domains (del Arco and Satrustegui, 1998; Kobayashi et al., 1999; del Arco et al., 2000; Palmieri et al., 2001; del Arco and Satrustegui, 2004). This subfamily contains two groups of proteins: the first subgroup consists of aralar1 and citrin, which are two isoforms of aspartate/glutamate carriers (del Arco and Satrustegui, 1998; Kobayashi et al., 1999). Aralar1 and citrin possess 678 and 675 amino acids respectively and are found in several tissues including brain (del Arco and Satrustegui, 1998; Kobayashi et al., 1999). The two proteins share 78% identity with each other. They both contain four EF‐hands (among them only three appear to be functional) in the N‐terminal domain, whereas the C‐terminal part possesses some identity with several mitochondrial protein carriers (> Table 27-1). Preliminary studies indicate that aralar1 binds calcium with high affinity (del Arco and Satrustegui, 1998). These proteins catalyze important steps in both the urea cycle and the aspartate/malate NADH shuttle (Palmieri et al., 2001). The second subgroup consists of the short calcium mitochondrial carriers (SCaMC) (del Arco and Satrustegui, 2004). Three human isoforms of SCaMC (about 480 amino acids) have been found in several tissues including brain (del Arco and Satrustegui, 2004). These proteins are expressed exclusively in mitochondria and share 70% identity with each other (del Arco and Satrustegui, 2004). They bind calcium with submicromolar affinity (Weber et al., 1997; Mashima et al., 2003). They have been suggested to play a role in nucleotide transport in mitochondria (del Arco and Satrustegui, 2004).

8

Conclusion

The regulatory EF‐hand CaBP or calcium sensors constitute a class of proteins with essential roles in the CNS. With an exponential increase in the number of targets, CaM continues to surprise with new targets such as the DAPK (Schumacher et al., 2004). CaM appears also to be the only calcium sensor precisely regulated at the gene level through its three different genes. The other families of calcium sensors (NCS, penta EF‐hand, CaBPs) seem more to be regulated via different protein isoforms such as the four KChIP family members regulating the A class potassium channel. The diversity of calcium sensor proteins, their complex tissue distribution, and their different calcium‐binding properties allow the various cells of the CNS to increase the diversity of intra‐ and extracellular signaling in response to complex calcium signals.

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Acknowledgments This work was supported in part by NCCR on Neural Plasticity and Repair and the Transregio Sonderforschungsbereich TR SFB 11 Konstanz/Zu¨rich.

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Calreticulin-Dependent Signaling During Embryonic Development

J. Groenendyk . M. Michalak

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 534 2 Calreticulin, a Ca2+-Buffering Chaperone of the ER . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 535 3 Calreticulin Outside the ER . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 536 4 Impaired Cardiogenesis in the Absence of Calreticulin (Loss-of-Function) . . . . . . . . . . . . . . . . . . . . . . 536 5 Calreticulin Signals Upstream of Calcineurin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 536 6 Calreticulin and Ca2+-Dependent Modulation of Transcriptional Pathways during Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 538 7 Gain-of-Function of Calreticulin and Embryogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 539 8 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 540

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Abstract: Ca2+ ions are signaling molecules responsible for controlling developmental processes, including fertilization, differentiation, proliferation, and transcription factor activation. Various cellular functions are regulated by changes in cytoplasmic Ca2+, including gene transcription and expression, protein synthesis, modification, folding and secretion, cell motility, cytoplasmic and mitochondrial energy metabolism, cell cycle progression, and apoptosis. Endoplasmic reticulum is an important Ca2+ storage organelle involved in virtually every aspect of Ca2+ homeostasis. Endoplasmic reticulum luminal Ca2+-binding chaperones such as calreticulin are critical for buffering endoplasmic reticulum Ca2+. In mice, calreticulin deficiency is lethal in utero because of the compromised Ca2+ storage capacity in the endoplasmic reticulum and disrupted InsP3 receptor-mediated Ca2+ release. A disturbance in Ca2+ release results in impaired cardiac development due to inhibited Ca2+-dependent transcriptional pathways. Calreticulin and the endoplasmic reticulum are the key upstream elements for calcineurin in Ca2+-signaling pathways. In contrast upregulation of calreticulin and overloading endoplasmic reticulum with Ca2+ leads to cardiac arrhythmias and impaired development of the cardiac conductive system. List of Abbreviations: DAG, diacylglycerol; HDAC, histone deacetylases; LRP1/CD91, LDL-receptor related protein; MEF2C, myocyte enhancer factor 2C; NF-AT, nuclear factor of activated T-cells; SERCA, sarcoplasmic/endoplasmic reticulum Ca2+-ATPase

1 Introduction Proper Ca2+ signaling is crucial from the earliest stages of development (Webb and Miller, 2003). For example, Ca2+ signaling plays a critical role during sperm invasion of the egg, with activation of the egg occurring by diffusion of a sperm-specific phospholipase C on sperm–egg fusion. Phospholipase C is responsible for cleaving the membrane phospholipid phosphatidylinositol bisphosphate (PIP2) into diacylglycerol (DAG) and inositol-1,4,5-trisphosphate (InsP3). DAG is retained at the plasma membrane, leading to activated protein kinase C, while InsP3 is a soluble molecule that traverses the cytoplasm, binds to the InsP3 receptor at the endoplasmic reticulum (ER) membrane and stimulates InsP3-mediated Ca2+ release from the lumen of the ER. The fertilization-triggered Ca2+ wave released from the ER controls cell cycle progression and cell division. Ca2+ signaling may be one of the very first signaling events to occur during embryogenesis (Malcuit et al., 2006; Whitaker, 2006). An important task of the fertilizationtriggered Ca2+ wave is to restart the cell cycle via Ca2+/calmodulin kinase II (CaMKII) pathways, dependent on Ca2+, that interact with the cell cycle control machinery. Other targets of Ca2+ signaling are protein kinase C, calcineurin, and the kinases/phosphatases of the cell cycle. Ca2+ signaling impacts the metaphase/ anaphase transition, and cytokinesis with separation of the daughter cells. In both oocytes and embryos, Ca2+ signals trigger a change in cell state and define the developmental program (Webb et al., 2005). Ca2+ signaling is ubiquitous, with significant fluctuations in cytoplasmic Ca2+ concentration serving as the trigger for the intracellular Ca2+-signaling pathway. For proper Ca2+ signaling to occur, the resting free cytoplasmic Ca2+ concentration has to be very low and maintained at a low level. Low levels are achieved by actively pumping Ca2+ from the cytoplasm into the ER or out of the cell as well as utilizing Ca2+-buffering proteins in the cytoplasm and in the lumen of the ER. Extracellular Ca2+ concentration is in excess of 2 mM, free cytoplasmic Ca2+ concentration is about 50–100 nM, while the free ER luminal Ca2+ concentration is roughly 200–300 mM, with total ER Ca2+ concentration up to 2–3 mM (Meldolesi and Pozzan, 1998). The ER, a major intracellular Ca2+ storage facility, maintains Ca2+ homeostasis within the cell by controlling Ca2+ release from the ER via the InsP3 receptor (InsP3R) and ryanodine receptor (RYR), with the ER stores being refilled by SERCA (sarcoplasmic/endoplasmic reticulum Ca2+-ATPase) (Berridge et al., 2003). In addition, Ca2+ is also pumped out of the cytoplasm into the extracellular space via the Na+/Ca2+ exchanger and the plasma membrane Ca2+-ATPase (PMCA) (Berridge et al., 2003). There is also a proficient coupling of intracellular Ca2+ release and extracellular Ca2+ entry mediated by a plasma membrane protein, Orail (Prakriya et al., 2006), and an ER transmembrane protein, Stiml (Liou et al., 2005;

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Huang et al., 2006). In the cytoplasm, Ca2+ exerts its effects by binding to Ca2+-regulated proteins such as calmodulin, a target for Ca2+ oscillations triggered upon Ca2+ signaling, as well as specific protein kinases and phosphatases which may modulate transcriptional processes involved in cell proliferation and differentiation and in organ growth and development. In this review, we focus on Ca2+ and ER luminal dynamics. Recent studies on the role of calreticulin, an ER-resident Ca2+-buffering chaperone, shed new light on the role of ER luminal Ca2+ during development.

2 Calreticulin, a Ca2+-Buffering Chaperone of the ER As the ER is an important source of intracellular Ca2+, Ca2+-buffering proteins within the ER, such as calreticulin, play an important role in cellular Ca2+ homeostasis (Michalak et al., 2002). Loss-of-function and gain-of-function of calreticulin have profound effects on Ca2+signaling (> Table 28-1). Calreticulin is

. Table 28-1 Consequences of calreticulin loss-of-function or gain-of-function on Ca2+ homeostasis Calreticulin deficiency Loss-of-function

Calreticulin upregulation Gain-of-function

Impact on Ca2+ homeostasis Inhibition of agonist-induced Ca2+ release (Mesaeli et al., 1999; Nakamura et al., 2001b; Guo et al., 2003; Martin et al., 2006) Reduced Ca2+ capacity of the ER (Nakamura et al., 2001b) Reduced free Ca2+ concentration in the ER lumen (Nakamura et al., 2001b) Delayed SOC influx (Mery et al., 1996; Nakamura et al., 2001b) Increased agonist induced Ca2+ release (Arnaudeau et al., 2002) Increased Ca2+ capacity of the ER (Xu et al., 2000; Arnaudeau et al., 2002) Increased free Ca2+ concentration in the ER lumen (Arnaudeau et al., 2002) Decreased SOC influx (Xu et al., 2000; Arnaudeau et al., 2002)

involved in the regulation of a number of functions specific to the ER, including Ca2+ storage and release (Mesaeli et al., 1999; Nakamura et al., 2001b), Ca2+ influx via SERCA (Camacho and Lechleiter, 1995), as well as store-operated Ca2+ entry at the plasma membrane (Arnaudeau et al., 2002). Another major role for calreticulin, as a member of the protein folding and quality control cycle located in the ER, is monitoring protein folding and trafficking (Michalak et al., 2002; Ellgaard and Frickel, 2003). Calreticulin binds to N-linked glycosylated proteins as well as nonglycosylated proteins (Ellgaard and Frickel, 2003). Specifically, the protein recognizes monoglucosylated carbohydrates on newly synthesized glycoproteins and, together with calnexin, plays a central role in quality control of the secretory pathway. The N-terminal domain and the central P-domain of calreticulin are responsible for chaperone (protein folding) activity (Nakamura et al., 2001b). In addition to its chaperone function, calreticulin is a major Ca2+-buffering ER luminal protein. The carboxyl-terminal region of calreticulin (C-domain) plays a key role in Ca2+-buffering/storage within the ER (Nakamura et al., 2001b). In the Ca2+ binding region of the protein, pairs of negatively charged amino acid residues coordinate the binding of Ca2+. The C-domain of calreticulin contains 19 pairs of acidic and negatively charged residues and binds over 50% of ER luminal Ca2+ with high capacity (25 mol of Ca2+ per mol of protein) (Baksh et al., 1992). Interestingly, the chaperone function of calreticulin is modulated by Ca2+ binding to the C-domain in response to agonist-induced fluctuating Ca2+ levels in the lumen of the ER. Calreticulin functions as an efficient chaperone when Ca2+ stores in the ER are full but its folding function is depressed when stores are depleted indicating that the ER Ca2+ homeostasis is tightly linked to protein synthesis, folding, and trafficking.

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3 Calreticulin Outside the ER Although calreticulin plays a crucial role in the lumen of the ER, the protein has been detected on the cell surface, where it may play a potential role in antigen-processing events and serve as a mediator of adhesion, as well as circulating outside the cell (Eggleton and Llewellyn, 1999). Cell surface calreticulin performs a distinct function as a receptor for thrombospondin, a cell surface ligand involved in focal adhesions and actin cytoskeleton (Goicoechea et al., 2000; Goicoechea et al., 2002), as well as interacting with LDLreceptor related protein (LRP1/CD91), a cell surface receptor (Ogden et al., 2001; Orr et al., 2003). T-lymphocyte adhesion to target cells performs a pivotal role in the function of the immune system, with thrombospondin interacting with LRP1/CD91 and calreticulin on the cell surface, regulating T-cell adhesion (Li et al., 2006). T-lymphocyte cells, part of the immune system, monitor the lymphatic system by migrating and interacting with endothelial cells and their extracellular matrix with frequent adhesive contacts. These interactions appear to regulate lymphocyte behavior and function and dictate cellular response, with thrombospondin performing an important role in conjunction with LRP/CD91 and calreticulin (Forslow et al., 2007). The interaction between thrombospondin and calreticulin may trigger T-lymphocyte motility and migration (Orr et al., 2003). A fascinating recent discovery indicates that calreticulin acts as a second generation recognition ligand at the cell surface upon an apoptotic signal, stimulating the LRP/CD91 on the surface of the engulfing cell (Gardai et al., 2005). Calreticulin plays an important role during integrin-mediated adhesion events (Fadel et al., 1999). Calreticulin is found on the cell surface of platelets and interacts with integrin a2b1 and glycoprotein VI, indicating a role for calreticulin in modulation of the platelet–collagen interaction (Elton et al., 2002). Calreticulin is implicated in signal transduction events during sperm–egg interactions at fertilization by interacting with egg cytoskeleton and mediating transmembrane signaling to the cell cycle (Tutuncu et al., 2004). The protein interacts with the DNA binding domain of the glucocorticoid and retinoic acid receptors (Burns et al., 1994) where it may facilitate Ca2+-dependent delivery of the receptor from the nucleus to the cytoplasm (Holaska et al., 2001). Recently in cancer cells, calreticulin is identified to translocate to the cell surface for recognition by T-cells and subsequent targeting for apoptosis upon treatment with anthracyclins (Obeid et al., 2007).

4 Impaired Cardiogenesis in the Absence of Calreticulin (Loss-of-Function) As calreticulin is 35% conserved across most species and given the role of calreticulin in a number of cellular process such as protein folding and Ca2+ homeostasis, it is not surprising that calreticulin deficiency is embryonic lethal in mice (Mesaeli et al., 1999). Cells derived from calreticulin-deficient embryos have impaired Ca2+ handling and compromised protein folding and quality control (Mesaeli et al., 1999; Nakamura et al., 2001b; Molinari et al., 2004). The major effects of calreticulin deficiency are observed in the heart (Mesaeli et al., 1999). Calreticulin-deficient embryos die at day 14.5 as a result of insufficient development of the ventricular wall (Mesaeli et al., 1999). Calreticulin-deficient embryonic stem (ES) cells also have inhibited cardiogenesis (Li et al., 2002; Mery et al., 2005). Both calreticulin-deficient mouse cardiomyocytes and ES cell-derived cardiomyocytes have disorganized myofibrils, indicating that calreticulin plays an important role during myofibrillogenesis (Li et al., 2002; Lozyk et al., 2006). Interestingly, these major differences are only detected in the developing ventricles while the atria of wild-type and calreticulin-deficient hearts are similar in appearance at all stages of development (Lozyk et al., 2006) (> Figure 28-1).

5 Calreticulin Signals Upstream of Calcineurin How does calreticulin contribute to Ca2+-dependent events during embryogenesis? Calreticulin is highly expressed in embryonic heart where it performs an important role by providing the Ca2+ necessary during cardiac growth and differentiation (Mesaeli et al., 1999). Studies on fibroblasts derived from calreticulin-deficient embryos indicate inhibition of agonist-mediated Ca2+ release from the ER in the

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. Figure 28‐1 Impact of loss-of-function and gain-of-function of calreticulin on development. Calreticulin deficiency leads to reduced Ca2+ concentration of the ER resulting in abnormal embryonic development, embryonic lethality and impaired myofibrillogenesis. Upregulation of expression of calreticulin increases Ca2+ capacity/content of the ER resulting in abnormal development of the cardiac conductive system

absence of calreticulin (Mesaeli et al., 1999; Nakamura et al., 2001b). Furthermore, the Ca2+ capacity of ER stores is significantly reduced in the absence of calreticulin, affecting proper function of the store-operated Ca2+ influx (Nakamura et al., 2001b). One target protein that is critically dependent on ER and cytoplasmic Ca2+ signaling is calcineurin, a Ca2+, calmodulin-dependent serine/threonine phosphatase. Calcineurin plays a vital role during cellular response to many diverse extracellular signals and stresses and is essential in the regulation of muscle differentiation, memory processes, and apoptosis (Crabtree and Olson, 2002). Calcineurin is a heterotetramer of A and B subunits. Calcineurin-A contains an N-terminal catalytic domain followed by a calmodulin binding site and an autoinhibitory (AI) region (Rusnak and Mertz, 2000). Calcineurin-B contains EF-hand Ca2+ binding motifs (Rusnak and Mertz, 2000) and Ca2+ binding to the protein promote the interaction between calcineurin-A and calcineurin-B to activate phosphatase activity (Stemmer and Klee, 1994). Calcineurin-A alone has very little endogenous phosphatase activity but phosphatase activity is significantly enhanced when it interacts with calcineurin-B. A constitutively active form of calcineurin-A (activated-calcineurin) can be generated by a C-terminal deletion of the AI domain and a portion of calmodulin-binding domain (Bandyopadhyay et al., 2000). Calcineurin regulates transcription factors responsible for alterations in gene transcription, in addition to affecting posttranslational modifications of proteins involved in Ca2+ homeostasis. Specifically, calcineurin controls gene expression via dephosphorylation of transcription factors, such as nuclear factor of activated T-cells (NF-AT), myocyte enhancer factor 2C (MEF2C), nuclear factor kB (NFkB), cyclic AMP response element-binding protein (CREB), and Elk-1, affecting their nuclear localization and ultimately is responsible for the up and/or downregulation of proteins involved in growth and development (Crabtree and Olson, 2002). Another target protein of Ca2+ and calmodulin is CaMKII. Upon Ca2+ and calmodulin

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binding, the multimeric kinase is released from autoinhibition and activated (Griffith, 2004). Downstream targets of CaMKII include cell cycle components, thereby playing a critical role during growth and development (Griffith, 2004). Cells isolated from calreticulin-deficient embryos have not only impaired agonist-induced Ca2+ release but also inhibited nuclear import of the transcription factor NF-AT (Mesaeli et al., 1999; Guo et al., 2002) and MEF2C (Lynch et al., 2005). Impairment in NF-AT import suggests that calreticulin may control availability of Ca2+ ions required for nuclear translocation of NF-AT during cardiac development. Indeed, activation of the NF-AT/GATA-4/calcineurin transcriptional pathway depends on Ca2+ release from the ER and Ca2+ translocation via plasma membrane Ca2+ channels (Crabtree and Olson, 2002). We hypothesized that calreticulin-deficient cells have inhibited calcineurin activity and, consequently, impairment in cardiac-specific transcriptional processes (Mesaeli et al., 1999). In support of this hypothesis, the lethality of calreticulin deficiency is rescued by the expression of constitutively active calcineurin in the heart (Guo et al., 2002), indicating that calreticulin is an essential molecule upstream from calcineurin during cardiac development. In the early stages of cardiac development, calreticulin is required to ensure normal Ca2+ release from the ER and thus proper activation of calcineurin and its associated transcriptional pathways (Lynch et al., 2006). Overexpression of activated-calcineurin renders calcineurin-dependent pathways less reliant on sustained elevation of cytoplasmic Ca2+ and therefore calcineurin-dependent pathways should be active even in the absence of Ca2+ release from the ER. These observations provide a molecular explanation for the embryonic lethality witnessed in calreticulin-deficient mice and indicate that calreticulin is a key upstream player in the Ca2+-signaling cascades which regulate calcineurin activity. It also highlights the importance of both calreticulin and calcineurin in Ca2+-dependent signaling cascades during early cardiac development. In the absence of calreticulin, expression of activated-calcineurin in the heart permits adequate progression of cardiac development during embryogenesis. Together, these findings show that calreticulin and calcineurin play important roles in the Ca2+-dependent pathways that are essential for myofibrillogenesis and normal cardiac development. As well, overexpression of calreticulin is also observed to promote differentiation-dependent apoptosis by suppressing the prosurvival kinase Akt, presumably through the release of Ca2+ and the activation of calcineurin (Kageyama et al., 2002) suggesting again a link between calreticulin and calcineurin (Lynch et al., 2006).

6 Calreticulin and Ca2+-Dependent Modulation of Transcriptional Pathways during Development The role of calcineurin in the NF-AT-signaling pathway in many tissues is well established (Crabtree, 2001). The NF-AT pathway is also critical in cardiac physiology and pathology (Frey et al., 2000). For example, NF-ATc-deficient mice die in utero due to impaired cardiac valves and septa development (delaPompa et al., 1998; Ranger et al., 1998). Overexpression of a constitutively active form of NF-AT or calcineurin in the heart causes severe cardiac hypertrophy (Molkentin et al., 1998). Do these pathways play a role in developing heart? The reversal of the embryonic lethality that results from calreticulin deficiency, by the expression of only one protein in the heart (activated-calcineurin), highlights the importance of both calreticulin and calcineurin in Ca2+-dependent signaling cascades during early cardiac development. The molecular details underlying the calcineurin-dependent rescue of lethality indicate that during cardiac development, calreticulin signals upstream of calcineurin, which in turn signal upstream from MEF2C (Lynch et al., 2005). MEF2 mediates a variety of cellular functions in neurons, skeletal muscle and the heart (Molkentin and Dorn, 2001; Frey and Olson, 2003). There are four genes encoding MEF2 proteins (MEF2A-D). MEF2C is highly expressed in developing heart and plays an important role during early cardiac development (Molkentin et al., 1996). MEF2 activity may be regulated by a variety of effectors in different cell types including mitogen-activated protein kinases (MAPKs) (p38 and BMK1/ERK5), Ca2+/calmodulindependent protein kinase and calcineurin (Wu et al., 2000). Cabin 1, MEF2-interacting transcriptional repressor (MITR) and the histone deacetylases HDAC4, 5, and 7 have been identified as MEF2-specific transcriptional repressors (Crabtree and Olson, 2002; Backs and Olson, 2006). In cardiomyocytes,

Calreticulin-dependent signaling during embryonic development

28

activated-CaMKIV disrupts the MEF2/HDAC complex and causes the unmasking of MEF2 transcriptional activity (Backs and Olson, 2006). In the cytoplasm, calmodulin can associate with the MEF2-binding domain of HDAC, perhaps representing a Ca2+-dependent feedback mechanism. With the HDACs removed, MEF2C activity can be enhanced by phosphorylation of the MEF2 transactivation domain by casein protein kinase II, protein kinase C (Black and Olson, 1998), and/or MAP-dependent protein kinase (Zhao et al., 1999). Additionally, HDAC removal permits recruitment of p300 by MEF2C and functions to amplify MEF2C transcriptional activity (Eckner et al., 1996). Recent studies indicate that in the developing heart, MEF2C is a target of calcineurin (Passier et al., 2000) and therefore is impacted by the ER and calreticulin. However, CaMKI and CaMKII are not effective in activation of MEF2C (Passier et al., 2000). Ca2+/calmodulin-dependent protein kinase and calcineurin pathways likely act in parallel and there must be important cross talk occurring between the two pathways.

7 Gain-of-Function of Calreticulin and Embryogenesis The conductive system of the heart is composed of the sinoatrial (SA) and the atrioventricular (AV) nodes. The SA nodes contain pacemaker cells, activated by an inward Ca2+ current, generating action potential which propagates along the conduction system and into the ventricles, producing contraction of the muscle. These pacemaker cells undergo spontaneous and repetitive depolarization, leading to a heart rhythm. These cells feature at their membrane a functional pacemaker channel, namely the hyperpolarization-activated cyclic nucleotide-gated K+ channel (HCN), generating the If current and setting the spontaneous and rhythmic action potentials in the heart. The molecular keystones of pacemaker activity in early stages of cardiogenesis are not understood. Spontaneous contractions are observed as early as embryonic day 7.5, even when the ion channel is not yet fully functional. These contractions are dependent on InsP3-induced intracellular Ca2+ signaling and Ca2+ capacity of the ER (Mery et al., 2005). Ca2+ signaling oscillates, generating Ca2+ waves involved in triggering activation of numerous Ca2+sensitive proteins, elicited by release of Ca2+ from the ER via the InsP3R (Mikoshiba, 2006). InsP3Rdependent release of Ca2+ is directly involved in providing a pacing mechanism for cardiomyocytes. Depolarization of pacemaker cells is triggered by InsP3-sensitive Ca2+ stores, demonstrated using pacemaker cardiomyocyte-differentiated ES cells that overexpress calreticulin, an ER localized Ca2+-buffering protein, or have knocked-down expression of the InsP3R (Mery et al., 2005), with excitability of these cells severely compromised. The calreticulin gene is highly active in the heart during early stages of development and is downregulated after birth (Mesaeli et al., 1999). Why is the calreticulin gene suppressed postnatally? High expression of calreticulin in postnatal heart results in congenital arrhythmia (bradycardia), sinus node depression, complete heart block, and death from heart failure (Nakamura et al., 2001a). As demonstrated, calreticulin has the ability to regulate differentiation and growth in a number of different manners, including regulation of intracellular Ca2+ as well as monitoring protein folding. Furthermore, phenotype of the calreticulin overexpressing mouse is reminiscent of complete heart block observed in children. The cause and molecular mechanisms involved in the complete heart block are not known. Yet, despite cardiac pacing, over 30% of these children die in the first year of life. Calreticulin must be part of a pathway responsible for the etiology of the disease and is necessary during growth and differentiation. Increased expression of calreticulin may affect the folding and function of Ca2+ channels (and other ion channels) and connexins/gap junctions (Nakamura et al., 2001a). A fascinating observation is that calreticulin overexpressing ES cell-derived embryoid bodies also exhibit problems with development of pacemaker activity (Mery et al., 2005). What function(s) of calreticulin are responsible for these unique phenotypes? Increased ER Ca2+ content in pacemaker cells over-expressing calreticulin prevents Ca2+-dependent generation of depolarizing currents in ES cell-derived cardiac pacemaker cells. These findings indicate that spontaneous action potentials do not depend on membrane potential but rather on intracellular Ca2+ stores (Mery et al., 2005) and provides the first evidence that ER-mediated Ca2+ dynamics and ER-associated Ca2+-buffering proteins are important in excitable cells at early stages of development. These observations also help explain severe cardiac

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arrhythmias, congenital heart block and sudden death in hearts with upregulated calreticulin and ER Ca2+ overload (Nakamura et al., 2001a; Mery et al., 2005). This evidence underscores the importance of the ER and Ca2+-dependent signaling pathways during the early stages of cellular commitment and cardiogenesis.

8 Conclusion The importance of Ca2+ signaling can be demonstrated by the numerous intracellular events influenced by disruptions in the signaling cascade, such as transcription factor activation, as well as cell–cell adhesion, directly affecting embryogenesis and development. Ca2+ signals are useful in a variety of developmental functions specifically related to cardiac function and neuronal development. As described in this review, these are just a few ways that calreticulin may regulate embryogenesis and development. Direct regulation of calreticulin expression is critical during embryogenesis with the Ca2+-buffering capacity of calreticulin responsible for the activation of a number of transcription factors involved in development. Calreticulin, as a Ca2+-buffering protein found in the lumen of the ER, appears to be a necessary component of Ca2+ regulatory systems in the developing embryo and influences Ca2+ and calcineurin-dependent pathways during embryogenesis.

Acknowledgments Work in our laboratory is supported by the CIHR and AHFMR. M.M. is a CIHR Senior Investigator.

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Voltage-Gated Calcium Channels

M. Wakamori . K. Imoto

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 544

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A Short History of VGCCs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 544

3

Nomenclature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 545

4 4.1 4.2 4.3 4.4 4.5

Molecular Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 545 a1 Subunit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 546 a2d Subunit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 547 b Subunit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 547 g Subunits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 547 Associated Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 548

5 CaV2.1 (P/Q-type) Calcium Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 548 5.1 Developmental Changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 549 5.2 P/Q Problem . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 549 6 CaV2.2 (N-type) Calcium Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 549 6.1 G-Protein Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 550 6.2 Developmental Change . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 550 7

R-Type Calcium Channel . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 551

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L-Type Calcium Channels and Gene Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 551

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T-type Calcium Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 552

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Role of VGCCS in Action Potential Generation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 552

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Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 553

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29

Voltage-gated calcium channels

Abstract: Voltage-gated calcium channels (VGCCs) are membrane proteins and mediate Ca2+ influx in response to membrane depolarization to evoke a wide spectrum of cellular responses, which include neurotransmitter release and activation of Ca2+-dependent enzymes. Molecularly, VGCCs are composed of multiple subunits, and their channel properties are primarily determined by the α1 subunits, which form the channel pore and various binding sites for associated proteins and drugs. There are ten genes encoding the α1 subunits. CaV2.1 (P/Q type) and CaV2.2 (N-type) are two major VGCCs in the brain, and are involved in neurotransmitter release. Recent studies revealed differences between them, for example in G-protein mediated modulation and in developmental changes. CaV1 (L-type) VGCCs are involved also in inducing changes in gene expression. CaV3 (T-type) VGCCs activate at subthreshold potentials, and for example play an important role in generating rhythmic activity. List of Abbreviations: AID, a1 interacting domain; CCAT, channel-associated transcriptional regulator; cDNA, complementary DNA; CGRP, calcitonin gene-related peptide; ChIs, cholinergic interneurons; GID, G-protein interaction domain; HVA, high-voltage activated; LVA, low-voltage activated; MSNs, medium spiny neurons; NALCN, sodium leak channel; NSCaTE, N-terminal spatial Ca2+ transforming element; PKC, protein kinase C; RIM1, Rab3-interacting molecule 1; TARP, transmembrane AMPA receptor regulatory protein; VGCC, voltage-gated calcium channels

1

Introduction

Voltage-gated calcium channels (VGCCs) are ion channels that are expressed widely and mainly in excitable cells, and mediate Ca2+ influx in response to membrane depolarization. Inward Ca2+ currents through VGCCs cause further membrane depolarization and in some cases generate Ca2+ spikes. Since Ca2+ functions as a critical intracellular messenger, Ca2+ influxes not only depolarize the membrane potential but also regulate various intracellular processes (Hille, 2001). Because of the dual roles of VGCCs controlling electrical and biochemical activities, VGCCs are involved directly and indirectly in a broad spectrum of cellular processes. VGCCs are abundantly expressed in neurons. VGCCs are an indispensable component of synaptic transmission. In presynaptic terminals, Ca2+ influx mediated by VGCCs in response to depolarization triggers the sequential events of neurotransmitter release. On the postsynaptic side, Ca2+ influx causes membrane depolarization, activation of Ca2+-dependent ion channels and enzymes. Ca2+ is also involved in the regulation of gene expression. Because of such important roles played by VGCCs, their activity is kept under control in many ways. Interactions with G proteins, calmodulin, and synaptic proteins are known to regulate channel activities. In addition, expression, transport, distribution, and turnover of VGCCs have to be tightly controlled, although the molecular mechanisms for such regulations are still mostly unknown. Excessive Ca2+ elevation (and probably the lack of sufficient Ca2+ influx, too) is detrimental to or catastrophic for many types of cells, and defects of VGCC regulations are often associated with neurological diseases.

2

A Short History of VGCCs

VGCC was first discovered in crab muscle fibers (Fatt and Ginsbor, 1958). Susumu Hagiwara and his coworkers made an extensive study of Ca2+ currents in a wide range of cells, which include invertebrate and vertebrate muscles, axons, nerve cells, and egg cells (Hagiwara and Byerly, 1981). The first step of classification of VGCCs was made on the basis of the voltage range of channel activation. High-voltageactivated (HVA) channels opened at more positive potentials (typically positive to 40 mV) than voltagegated sodium channels, whereas low-voltage activated (LVA) channels opened at more negative potentials (typically positive to 70 mV) (Hagiwara et al., 1975). The advent of the single-channel recording technique contributed to the refinement of the classification (Carbone and Lux, 1984). The HVA channels were named L-type, because of the long-lasting currents and the large single-channel conductance. The LVA channels were called T-type channels because of the transient currents and the tiny single-channel

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conductance. Dihydropyridines were the defining drug of L-type channels. It became clear soon, however, that neurons had dihydropyridine-insensitive HVA channels, which were then named N-type (for neuronal channels) (Nowycky et al., 1985). A peptide toxin o-conotoxin GVIA which was derived from Pacific cone snails Conus geographus, was found to block N-type channels specifically (Aosaki and Kasai, 1989). But there were yet other HVA channels. HVA channels in cerebellar Purkinje neurons were found insensitive to o-conotoxin GVIA, and were named P-type (for Purkinje cells) (Llina´s et al., 1989). Initial molecular cloning of VGCCs was conducted through a classical way, i.e., protein purification followed by amino acid sequencing and library screening. Shosaku Numa and his colleagues successfully cloned complementary DNA (cDNA) encoding the main subunit (a1) of the rabbit skeletal muscle VGCC (Tanabe et al., 1989). They purified the channel protein using photoaffinity-labeled dihydropyridine as a marker. Once a VGCC cDNA clone was obtained, it was used for low-stringency cross hybridization screening of cDNA libraries to yield cDNAs of other VGCC subtypes. When the first cDNA clone isolated from the brain was recombinantly expressed, the channel was insensitive to o-conotoxin GVIA (Mori et al., 1991). Since it was abundantly expressed in cerebellar Purkinje cells and was blocked by venom of a spider Agelenopsis aperta, later identified as o-agatoxin IVA, (Mintz et al., 1992), the recombinant channel was supposed to be P-type. But it was different from the native P-type channel of Purkinje cells in inactivation kinetics and sensitivity to o-agatoxin IVA. The recombinant channel was more similar to channels in cerebellar granule cells, and named Q-type (Wheeler et al., 1994). Calcium channel currents insensitive to dihydropyridines, o-conotoxin GVIA and o-agatoxin IVA were identified in cerebellar granule cells, and the VGCC was named R-type (residual) (Randall and Tsien, 1995). The F-type (CaV1.4) was identified through the genetic analysis of X-linked congenital stationary night blindness (Bech-Hansen et al., 1998). T-type VGCCs were difficult targets for molecular cloning, because they escaped cross hybridization screening. Edward Perez-Reyes and his colleagues smartly used the Genbank information and finally identified genes encoding T-type channels (Perez-Reyes et al., 1998).

3

Nomenclature

Although VGCCs are composed of multiple subunits (see the later section), the channel properties are primarily determined by the a1 subunits, which form the channel pore and various binding sites for associated proteins and drugs. It is now generally accepted that mammalians have 10 distinct genes encoding a1 subunits. The VGCC subtypes are molecularly named according to their a1 subunits (for example, VGCC containing the a12.1 subunit is CaV2.1 channel) (Ertel et al., 2000; Catterall et al., 2005). A number of studies in the 1990s contributed to relate the molecular VGCC classification to the classical classification based on electrophysiological and pharmacological properties (> Table 29-1). VGCCs are expressed not only in neurons but also in glial cells. Electrophysiological studies showed that rat cultured cortical astrocytes express functional CaV1, CaV2.2, and CaV2.3 calcium channels (D’Ascenzo et al., 2003). These channels are supposed to play substantial roles in the regulation of astrocyte Ca2+ influx and to influence glial functions and neuron-glia cross talks. In the rest of the review, neuronal calcium channels are discussed.

4

Molecular Structure

Biochemical properties of VGCCs were most thoroughly studied using skeletal muscle VGCC (CaV1.1). CaV1.1 channel is composed of a1, a2, b, g, and d subunits (Takahashi et al., 1987). Other HVA VGCCs are assumed to have similar molecular compositions, although the status of the g subunit is controversial. Since many VGCC subtypes are present in the brain, it is difficult to isolate a single VGCC subtype. A commonly used strategy to examine the subunit composition has been to coexpress subunits in various combinations. If the channel properties are significantly changed and the combined subunits are expressed in same tissues, the results are regarded as a strong evidence for the subunit combination.

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. Table 29-1 Classification of VGCCs

HVA

LVA

VGCC subtypes CaV1.1

Conventional names L, S

CaV1.2

L, C

CaV1.3

L, D

CaV1.4 CaV2.1 CaV2.2 CaV2.3 CaV3.1

L, F P/Q, A N, B R, E T, G

CaV3.2 CaV3.3

T, H T, I

Drugs Dihydropyridines, phenylalkylamines, and benzothiazepines

o-agatoxin IVA o-conotoxin GVIA SNX-482

Main tissues of expression Skeletal muscle Cardiac and smooth muscles Endocrine cells, nervous system Retina Nervous system

Cardiac muscle, nervous system Ni2+ Nervous system

In addition to various combinations of multisubunit assembly, another complicating factor is alternative RNA splicing. The a1, a2/d, b subunits are known to have alternative splice variants. Alternative splicing has significant influence on the channel properties. For example, exon 37 of the a12.2 subunit gene controls G protein-dependent inhibition of CaV2.2 channel (Raingo et al., 2007).

4.1 a1 Subunit The a1 subunit is the main and largest subunit of 190–250 kDa (~2,000 amino acid residues). It forms the conduction pore and various sites for channel regulation and drug interactions. The a1 subunit is organized in four homologous domains, like the a subunit of voltage-gated sodium channels. Each domain has six hydrophobic segments, which in a conventional model are considered to form transmembrane segments. The fourth segments (S4) have positively charged amino acid residues spaced regularly, and serve as the voltage sensor. A mutation that leads to a charge-neutralizing arginine-to-glycine substitution in S4 of repeat III expectedly reduced voltage sensitivity of activation in mutant mice, rolling Nagoya (Mori et al., 2000). VGCCs are selective for Ca2+ in a physiological condition. VGCCs are permeable to Ba2+ and Sr2+, but impermeable to Mg2+. Interestingly, Ca2+ must be present for the Ca2+ selectivity. In the absence of Ca2+, VGCCs become nonselective cation channels. This interesting property is explained by a model, which postulates the repulsion force between two Ca2+ at two Ca2+ binding sites in the selectivity filter (Almers and McCleskey, 1984; Hess and Tsien, 1984). The molecular correlate of the selective filter is four glutamic acid residues (one in each domain) located between S5 and S6 segments (Kim et al., 1993). Mutations of amino acid residues at the homologous locations are able to confer Ca2+ selectivity on voltage-gated sodium channels (Heinemann et al., 1992). The a1 subunits have binding sites for modulatory molecules. The a1-interacting domain (AID) of an 18-amino acid region located in the I-II linker is the binding site for the b subunits. AID forms an amphipathic helix in complex with the b subunit (Opatowsky et al., 2004). Binding with G-protein bg subunits is more complicated, because multiple regions within the a1 subunit contribute to form a binding pocket (De Waard et al., 2005). The determinants for the Gbg interaction include one region in the N-terminus, three in the I-II linker (the second partially overlaps AID; the third is called G-protein interaction domain (GID), and two in C-terminus. Sequences in the cytoplasmic II-III linker, termed synprint (synaptic protein interaction) site, play an important role in functional incorporation of VGCC into the synaptic vesicle fusion apparatus.

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The VGCC a1 subunits show sequence and structural similarities to the voltage-gated sodium channel a subunits. Besides the canonical voltage-gated calcium and sodium channels, a sodium leak channel (NALCN) shows sequence similarity (Lee et al., 1999; Lu et al., 2007). NALCN is voltage-independent and permeable to Na+, K+, and Ca2+. NALCN is expressed in all brain regions.

4.2 a2d Subunit The a2 and d subunits were identified independently (Ellis et al., 1988), but later they were shown to be the products of the same gene (De Jongh et al., 1990). The proproteins were cleaved posttranslationally. Four genes encoding a2d subunits have been cloned (a2d-1 to a2d-4). The a2 subunits are entirely extracellular, whereas the d subunits have a short transmembrane segment. The a2 subunit is extensively glycosylated. A mutation to make the proprotein insensitive to proteolytic cleavage reduced currents without affecting the voltage-dependence or kinetics of the channel (Andrade et al., 2007). The a2d subunit is known to bind gabapentin, a widely used anticonvulsant drug. Functions of the a2d subunit are extensively reviewed in Davies et al. (2007).

4.3 b Subunit The b subunits are cytoplasmic proteins of 50 kDa, and bind to an intracellular linker region of the a1 subunit. Four genes encode the b subunits (b1–b4). All b subunits show a number of splice variants. The skeletal muscles express the b1a subunit. The brain expresses b3, b4, b1b, and b2 subunits. Although there seems no strict association between a1 and b subunits, CaV2.2 channels more often associate with the b3 subunits. In cerebellar Purkinje cells, CaV2.1 channel associates with b2a and b4. The b subunit associates with the a1 subunit through the highly conserved, sodium leak channel (NALCN) in the a1 subunit. The b subunit increases the current amplitude and shifts the voltage dependence of activation. The b subunit aids in the trafficking of the a1 subunit to the plasma membrane, by masking an endoplasmic reticulum retention signal in the a1 subunit (Bichet et al., 2000). Besides the regulatory functions, the b subunits seem to play independent roles. Structural studies of the b subunits revealed that the b subunits have a PDZ-like, SH3 and guanylate kinase (GK) domains, and belong to the MAGUK family (Hanlon et al., 1999; Vendel et al., 2006). This observation suggests a role for the b subunits in scaffolding multiple signaling pathways around the channel. The b subunit is reviewed extensively in Dolphin (2003) and Hidalgo and Neely (2007).

4.4 g Subunits The g1 subunit, originally identified in skeletal muscle, is a protein of 32 kDa with four putative membranespanning regions. Eliminating the gene encoding the g1 subunit increased the calcium channel current and shifted the voltage dependence of inactivation to more positive membrane potentials measured in myotubes (Freise et al., 2000; Melzer et al., 2006). Genetic studies of stargazer mice with absence epilepsy led to the discovery of a protein named stargazin. Because of the sequence similarity to g1 subunit and the effects of coexpression on VGCC properties, stargazin was considered a neuronal VGCC g subunit (g2) (Letts et al., 1998). Subsequently several g subunits have been identified (up to g8). Stargazin (g2), g3, g4, and g8 function as transmembrane AMPA receptor regulatory protein (TARP) (Chen et al., 2000). g6, which shows the highest sequence similarity to g1, reduces cardiac CaV3.1 currents (Hansen et al., 2004). Coexpression of g7 abolishes CaV2.2 channel expression (Moss et al., 2002).

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4.5 Associated Proteins Calmodulin is a regulator across the CaV1–2 channels (Liang et al., 2003). Calmodulin associates constitutively with the C-terminal tail of the channels. Ca2+ binding of the C-terminal and N-terminal lobes of calmodulin induces distinct channel regulations. The C-lobe responds to local Ca2+ increases, whereas the N-lobe senses the global Ca2+ changes. In addition, the N-terminal Spatial Ca2+ Transforming Element of the channels (NSCaTE) confers a local spatial selectivity on the N-lobe (Dick et al., 2008). Neuronal VGCCs are modulated by the association with presynaptic SNARE proteins including syntaxin, SNAP-25 and synaptotagmin, primarily via interactions with the synprint region of a1 subunit. Syntaxin and SNAP-25 shifted steady-state inactivation curve of recombinant VGCCs to the hyperpolarizing direction by about 10 mV, indicating reduction of neuronal channel availability at presynaptic terminal (Bezprozvanny et al., 1995; Zhong et al., 1999). In contrast, formation of a mature SNARE complex concomitantly containing syntaxin, SNAP-25, and synaptotagmin, restored the inactivation curve (Zhong et al., 1999). This may suggest that VGCC associated with only one of the t-SNARE proteins cannot contribute to synaptic transmission, whereas VGCCs associated with the entire t-SNARE complement are capable of participating in vesicle release. The b subunits of VGCC complexes are also associated with the active zone protein Rab3-interacting molecule 1 (RIM1). The association of the RIM1 with the b subunit decelerated the speed of voltage-dependent inactivation and shifted steady-state inactivation curve of recombinant VGCCs to the depolarizing direction by about 30 mV (Kiyonaka et al., 2007). In addition to the anchoring of neurotransmitter-containing vesicles in the vicinity of VGCCs, the sustained Ca2+ influx by the RIM1-b interaction may contribute to neurotransmitter release through the inhibition of voltage-dependent inactivation.

5

CaV2.1 (P/Q-type) Calcium Channels

CaV2.1 (P/Q-type) and CaV2.2 (N-type) channels are probably most important VGCC subtypes for neurotransmitter release from nerve terminals. Analyses of neuromuscular junction showed that the number of packets of acetylcholine released is proportional to the fourth power of external Ca2+ concentration (Dodge and Rahamimoff, 1967). A similar relationship is considered to hold for CNS synapses. Based on the power relationship and several assumptions, contributions of each VGCC subtype to neurotransmitter release can be estimated. The assumptions include complete blockade of VGCCs with subtype-specific toxins and the same efficacy of Ca2+ influx through each VGCC subtype. In most of synapses, CaV2.1 and CaV2.2 channels are colocalized. But current components insensitive to o-agatoxin IVA, o-conotoxin GVIA, and L-type channel antagonists are also present. The insensitive component seems mainly composed of CaV2.3 (R-type) channel. At a synaptic level, CaV2 subtypes are located differently in calyx-type terminals, which are large presynaptic terminals and can be directly studied by patch-clamp (Wu et al., 1999). CaV2.1 channels trigger release more effectively than Ca2+ influx through CaV2.2 or CaV2.3 channels. A substantial fraction of CaV2.2 and CaV2.3 channels is located distant from release sites. It is still to be established whether CaV2 subtypes are differently distributed in presynaptic terminals of a usual small size. Since immunogold electron microscopy showed the most intense immunoreactivity for the a12.1 subunit was found in the presynaptic active zone of cerebellar parallel fiber varicosities (Kulik et al., 2004), CaV2.1 channel seems a more directly coupled VGCC subtype to mediate neurotransmitter release in a majority of small synapses. Another distinguishing feature of CaV2.1 is activity-dependent facilitation. During repetitive activation of the synaptic terminals, residual Ca2+ activates neuronal calcium sensor 1 and facilitates CaV2.1 currents (Tsujimoto et al., 2002). In mature mammalian neuromuscular junctions, CaV2.1 channel is the dominant subtype to mediate acetylcholine release (Uchitel et al., 1992; Rosato Siri and Uchitel, 1999). In some of neurons and synapses, one VGCC subtype dominates over others. CaV2.1 channel is the main subtype in cerebellar Purkinje cells (Llina´s et al., 1989). Therefore, it is expected that the synapses from Purkinje cells to deep cerebellar nuclei are mainly dependent on CaV2.1 channel. Some inhibitory

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interneurons show distinct preference of CaV2.1 and CaV2.2 channels. Cultured hippocampal inhibitory interneurons in stratum lucidum and stratum oriens use CaV2.1 channels for neurotransmitter release, whereas interneurons in stratum radiatum use CaV2.2 channels (Poncer et al., 1997). In hippocampal slice preparations, CaV2.1 channels mediate release at parvalbumin-expressing interneuron synapses, whereas CaV2.2 channels trigger synchronous and asynchronous release in cholecystokinin-expressing interneuron synapses (Hefft and Jonas, 2005). Such a preferential dependency on VGCC subtypes is also observed in prefrontal cortex. CaV2.1 channels mediate GABA release from fast-spiking interneurons (Zaitsev et al., 2007). Patchwork distribution of CaV2 channels may therefore be important for terminal-specific modulation of transmitter release (Reid et al., 2003).

5.1 Developmental Changes Whereas CaV2.2 channels are responsible for the majority of excitatory transmitter release early in development, CaV2.1 channels become more prominent during maturation in most systems studied (Iwasaki and Takahashi, 1998; Iwasaki et al., 2000). Because of the delay of CaV2.1 channel expression, CaV2.1 channel knockout mice are not embryonic lethal, but develop a rapidly progressive neurological deficit with specific characteristics of ataxia and dystonia before dying 3–4 weeks after birth (Jun et al., 1999). CaV2.1 mutant mice tottering start showing ataxia and epilepsy at 3 weeks of age (Noebels and Sidman, 1979; Fletcher et al., 1996).

5.2 P/Q Problem After molecular cloning and recombinant expression of CaV2.1 channel, there was a controversy regarding the relation between slow-inactivating P-type and fast-inactivating Q-type channels. It is now generally accepted that the main a1 subunits of the P- and Q-type channels are gene products of the single gene. Differences in subunit combination (particularly b subunit) and alternative splicing produce the different channel properties. While the b1b and b3 subunits caused a significant acceleration of inactivation kinetics, the b2a subunit dramatically slowed the inactivation (Stea et al., 1994). Asp-Val insertion (located between S3 and S4 of domain IV) caused by an alternative splicing slows inactivation (Bourinet et al., 1999). On the other hand, however, splice variants isolated from Purkinje cells failed to generate P-type channels (Tunemi et al., 2002). Posttranslational processing or modification by interacting proteins seems required for generating P-type currents.

6

CaV2.2 (N-type) Calcium Channels

Originally, the decaying component of high-voltage-activated (HVA) Ca2+ channel was assigned to N-type channel (Nowykey et al., 1985), but a wide variety of inactivation properties was reported (Aosaki and Kasai, 1989; Jones and Marks, 1989; Plummer and Hess, 1991). N-type channel was distinguished from other types in neurons by o-conotoxin GVIA-sensitivity. The CaV2.2 (N-type) is another major VGCC in the brain and is involved in neurotransmitter release at nerve terminals. Since it was the only identified neuronal VGCC until CaV2.1 was discovered, it was rather surprising to find that CaV2.2 knock-out mice were almost normal (Ino et al., 2001). Although o-conotoxin GVIA-sensitive CaV2.2 Ca2+ channel current was selectively eliminated in cell body of superior cervical ganglion (SCG) and dorsal root ganglion (DRG) neurons dissociated from 8 week-old CaV2.2-knock-out mice without compensations by other types of VGCCs (Ino et al., 2001; Hatakeyama et al., 2001), other types can substitute for CaV2.2 channel at the early stage of development, especially at synapses. Examinations of the functional roles of CaV2.2 Ca2+ channel indicate a pathophysiological role in the pain pathway in the spinal cord. First, CaV2.2 channels were located on the superficial laminae of the dorsal horn where primary afferent nerves terminate (Kerr et al., 1988). Second, CaV2.2 channel currents were

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detected in small sensory neurons of DRG, which are responsible for the mediation of nociceptive stimuli, as determined by the whole-cell patch-clamp technique (Scroggs and Fox 1989). Third, the specific CaV2.2 channel antagonist, o-conotoxin GVIA inhibited the release of calcitonin gene-related peptide (CGRP) and substance P from primary afferent nerves (Holz et al., 1988; Maggi et al., 1990; Santicioli et al., 1992). Fourth, the selective block of CaV2.2 channels via intrathecal administration of o-conotoxin GVIA or oconotoxin MVIIA significantly depressed the formalin phase 2 response, thermal hyperalgesia, mechanical allodynia, and postsurgical pain (Vanegas and Schaible, 2000). Finally, CaV2.2-knock-out mice exhibited diminution in the formalin phase 2 nociceptive response and thermal nociceptive response in the hot plate test (Hatakeyama et al., 2001). CaV2.2 Ca2+ channel is a therapeutic target of cardiovascular diseases as well as L-type channel. A unique dihydropyridine derivative, cilnidipine, has a dual antagonistic action on both L- and CaV2.2 channels. It inhibited the excessive release of catecholamine at the time of blood pressure reduction (Konda et al., 2005), thereby exhibiting a gradual, strong, and persistent antihypertensive effect without causing reflex tachycardia (Fujii et al., 1997).

6.1 G-Protein Inhibition Dunlap and Fischbach first reported the rapid inhibition of Ca2+ currents in chick DRG neurons by noradrenaline and serotonin (Dunlap and Fischbach 1981). Many hormones and neurotransmitters, acting via seven transmembrane G-protein-coupled receptors, modulate native neuronal Ca2+ currents. In rat sympathetic neurons where the main Ca2+ channel is CaV2.2 channel, five different modulatory pathways are used by 10 transmitters that inhibit Ca2+ current (Hille, 1994). The CaV2 channels are modulated through activation of G protein-coupled receptors. Among three CaV2 channels, CaV2.2 is the most sensitive to G-protein-mediated inhibition. The modulation mediated by Gbg subunits includes (1) inhibition of current amplitude, (2) slowing of current activation, (3) depolarizing shift of the voltage dependence of activation, and (4) relief of inhibition by strong depolarization (Ikeda, 1996; Herlitze et al., 1996). These phenomena are explained by the simple model that Gbg shifts the Ca2+ channels from ‘‘willing’’ to ‘‘reluctant’’ gating modes (Bean, 1989; Elmslie et al., 1990). Each gating mode has closed and open states, and the midpoint voltages of channel activation were 15 and 62 mV for the ‘‘willing’’ and the ‘‘reluctant’’ modes, respectively (Bean, 1989). At weak depolarizations, the reluctant closed state (RC) reluctant open state (RO) equilibrium favors RC, and the mode shift from the RC to the willing close state (WC) dominates, producing small current amplitude and slow current activation. However, at strong depolarizations RO is favored over RC, so the transition from RC to RO can occur first and the inhibition can be relieved. Recently, a new kinetic model which incorporates G protein dissociation was proposed (Weiss et al., 2006).

6.2 Developmental Change During aging from 14 to 270 days, total HVA Ca2+ current recorded from both medium spiny neurons (MSNs) and cholinergic interneurons (ChIs) of mouse striatum was unchanged. However, CaV2.2 component decreased from 36% at 14-day-old to 23% at 23-day-old in MSNs, and from 39% at 23-day-old to 15% at 40-day-old in ChIs, remaining constant before and thereafter. L-type component also exhibited the same developmental decrease, whereas CaV2.1 and CaV2.3 components showed a tendency to increase. Differently from striatal neurons, CaV2.2 component did not show a similar rearrangement in pyramidal neurons dissociated from cortical layers IV-V (Martella et al., 2008). Also, at thalamic and cerebellar synapses, the contribution of CaV2.2 Ca2+ channels to synaptic transmission is limited to the early postnatal stage and CaV2.1 channels play a predominant role in mature synaptic transmission (Iwasaki et al., 2000). These results show that distribution of Ca2+ channels is reconfigured during development.

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7

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R-Type Calcium Channel

CaV2.3 is widely distributed not only in the peripheral nervous system and CNS, but also in the endocrine (Jing et al., 2005), cardiovascular (Lu et al., 2004), sperm (Westenbroek and Babcock, 1999), and gastrointestinal system (Grabsch et al., 1999). Compared with CaV2.1 andCaV2.2 calcium channels, the functional role of CaV2.3 channel is less extensively studied. CaV2.3 channel shares some similarities with LVA, T-type Ca2+ channel, in the aspects of (1) high sensitivity to Ni2+, (2) rapid current inactivation, and (3) voltagedependence of inactivation (Ellinor et al., 1993; Soong et al., 1993; Wakamori et al., 1994). But although CaV2.3 channel is a member of HVA channels, CaV2.3 channel is activated at high-voltage range. Its singlechannel conductance 12–14 pS, (Ellinor et al., 1993; Wakamori et al., 1994) is almost the same as those of CaV2.1 13–17 pS, (Mori et al., 1991; Wakamori et al., 1998) and CaV2.2 channels 14 pS, (Fujita et al., 1993). From a practical point of view, CaV2.3 channel is defined by its resistance to dihydropyridines, o-conotoxin GVIA, and o-agatoxin IVA, but can be blocked by SNX-482, a peptide toxin isolated from tarantula venom (Newcomb et al., 1998). The SNX-482-sensitive portion was absent from CaV2.3 knock-out mice, whereas the size of SNX-482-resistant residual currents was not reduced (Wilson et al., 2000). Generation of CaV2.3-knock-out mice has provided detailed insight into the functional relevance of this channel. Within the CNS, CaV2.3 is involved in neurotransmitter release, spatial memory, and mossy fiber long-term-potentiation (Kubota et al., 2001; Breustedt et al., 2003; Dietrich et al., 2003). In addition, CaV2.3 is reported to be involved in control of pain behavior (Saegusa et al., 2000), myelinogenesis (Chen et al., 2000), fear-related emotion (Lee et al., 2002), and epileptogenesis (Weiergraber et al., 2006). Furthermore, CaV2.3 channel plays a protective role in ischemic neuronal injury (Toriyama et al., 2002). Except nervous system, CaV2.3 is essential for hormone secretion and glucose homeostasis (Pereverzev et al., 2002), and for impulse generation and conduction in heart (Weiergraber et al., 2005). CaV2.3 currents have been shown to participate in the formation of afterdepolarizations, plateau potentials, and bursting activity in hippocampal CA1 pyramidal neurons (Kuzmiski et al., 2005; Metz et al., 2005). Studies also indicate that native CaV2.3 channel mediates Ca2+ entry into dendritic spines (Sabatini and Svoboda, 2000). Influxed Ca2+ induces Ca2+ dependent inactivation in L-type Ca2+ channel (Eckert and Chad, 1984), whereas, surprisingly, Ca2+ influx through CaV2.3 channels has opposite effects on the channel activity itself. At lower cytosolic Ca2+ concentrations, a positive feedback mechanism, which includes activation through protein kinase C (PKC), increased in current amplitude, slowed down inactivation and speeded up recovery from short-term inactivation (Leroy et al., 2003; Klo¨ckner et al., 2004; Dietrich et al. 2003) proposed that CaV2.3 contributes selectively to the residual [Ca2+]i which also underlies various forms of synaptic plasticity but contributes less to neurotransmitter release. The initial positive feedback mechanisms based on PKC activity might later be attenuated by N-lobe calmodulin-dependent negative feedback loops (Liang et al., 2003), and therefore help to maintain physiological [Ca2+]i.

8

L-Type Calcium Channels and Gene Expression

L-type calcium channels are main calcium channels in skeletal muscles, cardiac muscles, and endocrine organs, but they are also present in neurons. The principal location of actions of L-type channels is postsynaptic (Ahlijanian et al., 1990). L-type calcium channels seem particularly effective at inducing changes in gene expression that underlie plasticity, adaptive neuronal responses, and survival (Bading et al., 1993). Calcium influx through L-type calcium channels activates transcription factors such as CREB, MEF, and NFAT that leads to the expression of genes such as c-fos and BDNF. There are several mechanisms that link L-type calcium channels to the activation of transcription factors. Ca2+ entering through calcium channels can diffuse to the nucleus and activate nuclear calcium-dependent enzymes, such as CaMKIV. Ca2+ can activate signaling cascades locally around the inner channel mouth, which propagate the signal to the nucleus (Dolmetsch et al., 2001). A recently reported mechanism is that the C-terminus of CaV1.2 channel, named Calcium

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Channel-Associated Transcriptional regulator (CCAT), is cleaved (Gomez-Ospina et al., 2006). C-terminal cleavage is also reported for CaV1.3 and CaV2.1 channels. Tumor suppressor elF3e is involved in internalization of CaV1.2 (Green et al., 2007).

9

T-type Calcium Channels

Molecular aspects of T-type VGCCa are reviewed in Peres-Reyes (2006). T-type calcium channels are encoded by three distinctive genes. Heterologous expression studies using HEK-293 cells demonstrated similarities and differences among three members of T-type calcium channels (Klo¨ckner et al., 1999). The three members shared the voltage-dependence of channel availability (steady-state inactivation). CaV3.1 showed the fastest activation and inactivation kinetics, as well as fast recovery from short-term inactivation. On the hand, CaV3.3 showed the fastest deactivation and the fastest recovery from long-term inactivation, whereas CaV3.2 showed the slowest recovery from long-terms inactivation. T-type calcium channel has been famous for its nickel sensitivity. But CaV3.2 is the only nickel-sensitive subtype (IC50 for block = 5–10 mM), whereas CaV3.1 and CaV3.3 are as nickel-insensitive as most high-voltage-activated calcium channels (IC50 for block 100 mM). The molecular determinant of nickel inhibition is the histidine residue (H191) located extracellularly in the S3-S4 loop of domain I (Kang et al., 2006). In contrast to high-voltage-activated calcium channels, the expression of a CaV3.x a1 subunit alone produced large currents. Studies of coexpression with b subunits or a2d subunits have been inconclusive. It has remained an open question whether there are any T-type channel auxiliary subunits. T-type calcium channels, particularly CaV3.1, are blocked by a scorpion peptide toxin kurtoxin (Chuang et al., 1998). Kurtoxin also reduces HVA channel currents (Sidach et al., 2002). In brain, all the three types are expressed. A high expression of CaV3.1 was observed in cerebellar Purkinje cells and granule cells. CaV3.3 is also expressed in them (Talley et al., 1999). Thalamic relay neurons are one of the well-documented neurons where T-type calcium channels play an important role in generating rhythmic activities (Jahnsen and Llina´s, 1984). In the rat thalamus, CaV3.1 is found predominantly in relay neurons, and thalamic reticular nucleus neurons strongly express CaV3.3 and moderately CaV3.2. A similar expression pattern is shown in the thalamus of primates (cynomolgus macaque). T-type VGCCs are modulated by hormones and neurotransmitters, mainly through activation of G-protein coupled receptors. Three T-type subtypes show differential patterns of G-protein coupling. For example, M1 muscarinic receptor activation leads to a strong inhibition of CaV3.3 through Gaq/11, but there is no effect or a moderate stimulation effect on CaV3.1 and CaV3.2 (Hildebrand et al., 2007). CaV3.1 and CaV3.3 are also regulated by Rho-associated kinase (ROCK) (Iftinca et al., 2007). CaV3.2 calcium channel is also regulated by extracellular Zn2+. Chelating Zn2+ with L-cysteine or the application of reducing agents enhances T-type calcium channel currents and lowers the threshold for excitability in C-type dorsal root ganglion neurons. T-type VGCCs are extensively reviewed by Nilius et al. (2006).

10

Role of VGCCS in Action Potential Generation

In cardiac muscles, action potential is generated by voltage-gated sodium channels, but the plateau phase of a long duration is maintained by Ca2+ influx mediated by VGCCs. In contrast, action potentials in neurons are usually very brief. The rising phase of the action potentials is generated by voltage-gated sodium channels, and HVA VGCCs become fully activated in the falling phase of action potentials. Since blocking VGCCs often results in a broadening of action potentials, VGCCs in neurons do not take part in generating action potentials but are more engaged in repolarization (Bean 2007). This function is based on the tight functional and structural association of VGCCs with large conductance calcium-activated potassium channels (BK channels) (Berkfeld et al., 2006). A simulation study suggests that HVA currents are much smaller than T-type currents in bursting CA3 pyramidal neurons (Xu and Clancy, 2008).

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Calcium currents are more important in generating action potentials at subthreshold potentials. Roles of subthreshold calcium currents in spontaneous firing were quantitatively analyzed in acutely dissociated midbrain dopaminergic neurons (Puopolo et al., 2007). Since dissociated neurons had lost most of the dendritic tree, the properties studied correspond to those of the somata. The application of o-agatoxin IVA, but not of o-conotoxin GVIA, slowed pacemaking. In some of the neurons, spontaneous firing persisted in the presence of tetrodotoxin (TTX), although spontaneous activity in TTX had broader and smaller spikes than in control condition. Ca2+ currents were larger than sodium currents at interspike voltages from 70 to 50 mV, whereas sodium currents were larger at potentials positive to 45 mV. Therefore, HVA calcium currents play significant roles in generating action potentials, at least, in some types of neurons. VGCCs are present in dendrites. In proximal apical dendrites of layer V pyramidal neurons, action potential evoked calcium transients are mediated by VGCC subtypes, which include not only CaV2.1 and CaV2.2 but also CaV2.3 and CaV1.3 channels (Markrum et al., 1995). In more distal portions of apical dendrites of layer V pyramidal neurons, bursts of back-propagating action potentials above the critical frequency of 10–20 Hz, elicited large regenerative potentials, which led to further depolarization at the soma (Larkum et al., 1999).

11

Conclusions

In the last two decades, we made a great advance in elucidating the function of VGCCs. For example, the outline role of VGCCs in neurotransmitter release is now mostly known. But there are still many unsolved problems. In future, it will be particularly important to study the functional role of somatic and dendritic VGCCs in processing information in the brain.

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Neural Roles of CLC Chloride Channels

S. Uchida . S. Sasaki

1 1.1 1.2 1.3 1.4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 560 ClC-1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 560 ClC-2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 560 ClC-3 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 560 ClC-6 and ClC-7 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 561

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Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 562

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Neural roles of CLC chloride channels

Abstract: Neural roles CLC chloride channels have been clarified by finding the mutations of human CLC genes in certain human diseases and by generating the CLC chloride channel knockout mice. Since five of the nice CLC channels are (CIC-3 through CIC-7) localized in membranes of intracellular organelles, studies on these channels are important for the better understanding of the roles of vesicular anion transport on various neural functions. List of Abbreviation: NCL, neuronal ceroid lipofuscinosis

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Introduction

The CLC chloride channel family consists of nine members in mammals, all of which are expressed in the central and/or peripheral nervous systems (Uchida, 2000; Jentsch et al., 2002; Uchida et al., 2005). Mutations in CLC genes in human inherited diseases have been found in the CLCN1 (Steinmeyer et al., 1991; Koch et al., 1992), CLCN2 (Haug et al., 2003), CLCN5 (Lloyd et al., 1996), CLCN7 (Kornak et al., 2001), and CLCNKB (Simon et al., 1997) genes. Apparent neural abnormalities were also found in the Clcn2 (Bosl et al., 2001), Clcn3 (Stobrawa et al., 2001; Yoshikawa et al., 2002), Clcn6 (Poet et al., 2006), and Clcn7 (Kornak et al., 2001; Lange et al., 2006) knockout mice. These results clearly suggest that CLC chloride channels have important roles in neural functions. This review focuses on the neural role of intracellular CLC chloride channels (ClC-3, ClC-4, ClC-6, and ClC-7) and also briefly describes the roles of ClC-1 and ClC-2, based on observations of CLC channel knockout mice.

1.1 ClC-1 ClC-1 is a skeletal muscle–specific ClC chloride channel. It constitutes dominant plasma membrane conductance of skeletal muscle. Accordingly, loss of function of ClC-1 in skeletal muscle leads to increased muscle excitability. Mutations in the human CLCN1 gene were found in patients with myotonia (Steinmeyer et al., 1991; Koch et al., 1992), and mutations in the mouse Clcn1 gene (Steinmeyer et al., 1991) were found in myotonic mice. These findings confirmed the physiological role of ClC-1 in plasma membrane conductance in skeletal muscles. Numerous mutations in the CLCN1 gene have been found, and these naturally occurring mutations have clarified the functionally important domains in ClC channels (for details, see review by Pusch, 2002).

1.2 ClC-2 ClC-2 is ubiquitously expressed in plasma membranes (Thiemann et al., 1992). Initially, it was postulated to be a volume-regulated chloride channel; however, its functional property as an inwardly rectifying chloride channel was different from that of a well-known volume-regulated chloride current. Clcn2 knockout mice were generated by the Jentsch group in 2001 (Bosl et al., 2001). These mice experience progressive retinal degeneration, but the mechanism of cell death in the retina remains unclear. In 2003 (Haug et al., 2003), mutations in the human CLCN2 gene were found in patients with idiopathic generalized epilepsies. The reason for the discrepancy between the human and mouse phenotypes remains to be elucidated.

1.3 ClC-3 ClC-3 (Kawasaki et al., 1994, 1995) forms a branch of the CLC family with ClC-4 and ClC-5 (Sakamoto et al., 1996). Unlike the predominant expression of ClC-5 in the kidney, ClC-3 and ClC-4 are abundantly expressed in the brain and other nervous system tissues. ClC-3 is localized in endosomes and synaptic vesicles. The disruption of ClC-3 in mice leads to developmental retardation, blindness, motor coordination deficit, and spontaneous hyperlocomotion (Stobrawa et al., 2001; Yoshikawa et al., 2002). Histological

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analysis of tissue from Clcn3 knockout mice revealed progressive degeneration of the retina, hippocampus, and ileal mucosa (Yoshikawa et al., 2002), which resembled the phenotypes observed in cathepsin D knockout mice. Cathepsin D deficiency causes lysosomal accumulation of ceroid lipofuscin containing the mitochondrial F1F0ATPase subunit c. In the Clcn3 knockout mice, subunit c accumulated heavily in the lysosomes (Yoshikawa et al., 2002). Accordingly, the Clcn3 knockout mouse is an animal model of a human lysosomal storage disease, neuronal ceroid lipofuscinosis (NCL). NCL may be caused by impaired endosomal acidification since ClC-3 is important for maintaining intra-vesicular pH by shunting proton pump currents (Hara-Chikuma et al., 2005). This finding was verified by the direct measurement of intra-vesicular pH. Recently, however, it was found that ClC-4 and ClC-5 are not chloride channels; rather, they are Cl/H antiporters (Scheel et al., 2005). This mode of Cl transport may not be beneficial for lowering intra-vesicular pH. Accordingly, ClC-3 may function primarily as a Cl uptake system rather than a shunt for proton currents. The role of intra-vesicular chloride in protein degradation systems must be investigated to elucidate the physiological role of ClC-3. Although a similar neuronal role for ClC-4 is anticipated, there has been no report on Clcn4 knockout mice.

1.4 ClC-6 and ClC-7 ClC-6 and ClC-7 form another branch in the CLC family. Like ClC-3, ClC-4, and ClC-5, both ClC-6 and ClC-7 reside in intracellular organelles (Suzuki et al., 2006). ClC-6 and ClC-7 are expressed in many tissues including the brain (Kida et al., 2001). It is unclear whether ClC-6 and ClC-7 function as Cl channels or Cl/H antiporters. In 2001, ClC-7 knockout mice were generated (Kornak et al., 2001). Although ClC-7 were thought to be ubiquitously expressed, the analysis of Clcn7 knockout mice, in which the lacZ gene replaced exons 7–10 and was fused in frame with exon 6, revealed that ClC-7 was abundantly expressed in osteoclasts. Although osteoclasts are present in normal numbers, they fail to resorb bone because they cannot acidify the extracellular resorption lacuna. ClC-7 is highly expressed in the ruffled membranes of osteoclasts, suggesting that ClC-7 is also required for efficient proton pumping by H-ATPase, like ClC-3 and ClC-5 in endosomes. This knockout phenotype has also been found in humans (Kornak et al., 2001). Patients with infantile malignant osteopetrosis have mutations in the CLCN7 gene. In addition to the phenotype in bone, ClC-7 knockout mice showed retinal and hippocampal degeneration. Following the initial observation by Uchida’s group that the disruption of one of the intracellular CLCs, ClC-3, in mice leads a phenotype similar to human NCL; Jentsch’s group investigated this possibility in Clcn7 knockout mice (Kasper et al., 2005). They found that CNS degeneration in Clcn7 knockout mice displayed features of NCL: electron-dense, osmiophilic material in the perikarya of hippocampal and cortical neurons that contained subunit c of F1F0 ATPase and autofluorescent pigment in the CA3 regions of the hippocampus. According to their analysis, these NCL phenotypes were much more severe in Clcn7 knockout mice than in their own Clcn3 knockout mice (Stobrawa et al., 2001), although Yoshikawa et al. clearly demonstrated NCL phenotypes in the Clcn3 knockout mice (Yoshikawa et al., 2002). The reason for this difference remains unclear and may be due to different genetic backgrounds and/or targeting strategies. However, the lysosomal pH in the Clcn7 knockout mice was not significantly reduced compared with that in wild-type mice. This observation raises the question of whether intravascular pH regulation is the main function of intra-vesicular CLC chloride channels (> Figure 30‐1). Recently, the generation and analysis of Clcn6 knockout mice was also reported by Jentsch’s group (Poet et al., 2006). Immunofluorescence analysis revealed that ClC-6 was present in punctate structures in neuronal somata and that it co-localized with lamp1. Fractional centrifugation using Percoll revealed that the intracellular localization of ClC-6 is closer to that of ClC-7 than ClC-3. They concluded that ClC-6 is predominantly expressed in late endosomes. Analysis of Clcn6 knockout mice revealed that neurons in almost all brain regions showed autofluorescence. A typical feature of NCL, the accumulation of saposin D and/or subunit c of the mitochondrial ATPase, was confirmed in the axon initial segments of Clcn6 knockout mice. This finding is consistent with the view, first pointed out by Uchida’s group, that the disruption of vesicular CLC leads to a phenotype similar to human NCL. However, despite these pathological changes, there were no obvious signs of neuronal death or degradation. Only impaired nociception and

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Neural roles of CLC chloride channels

. Figure 30‐1 Vesicular CLC chloride channels. In a heterologous expression system (Suzuki, et al. 2006), ClC-3, ClC-4, and ClC-5 showed a high degree of co-localization, but also showed some overlap with ClC-6 and ClC-7. Each CLC channel may not be strictly compartmentalized in native cells as shown in the figure

mild behavioral abnormalities were observed in Clcn6 knockout mice. As observed in the Clcn7 knockout mice, the lysosomal pH was not changed in the Clcn6 knockout mice.

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Summary

Five of the nine members of the CLC chloride channel family turned out to be vesicular chloride channels. The genes for two members (ClC-5 and ClC-7) are responsible for human diseases. Based on knockout mouse studies, the other members also have important roles in neuronal functions. Initially, these vesicular chloride channels were thought to simply provide a Cl shunt pathway for the efficient pumping of H-ATPase. However, ClC-4, ClC-5, and possibly ClC-3, have turned out to be antiporters rather than chloride channels. This finding suggests a new notion that intravesicular chloride, rather than pH, may be of primary importance for proper protein degradation in lysosomes. Further studies on this issue are exciting and may be important not only for neuronal cell biology, but also for general cell biology.

References Bosl MR, Stein V, Hubner C, Zdebik AA, Jordt SE, et al. 2001. Male germ cells and photoreceptors, both dependent on close cell-cell interactions, degenerate upon ClC-2 Cl(-) channel disruption. EMBO J 20: 1289-1299. Hara-Chikuma M, Yang B, Sonawane ND, Sasaki S, Uchida S, et al. 2005. ClC-3 chloride channels facilitate endosomal acidification and chloride accumulation. J Biol Chem 280: 1241-1247. Haug K, Warnstedt M, Alekov AK, Sander T, Ramirez A, et al. 2003. Mutations in CLCN2 encoding a voltage-gated chloride channel are associated with idiopathic generalized epilepsies. Nat Genet 33: 527-532. Jentsch TJ, Stein V, Weinreich F, Zdebik AA. 2002. Molecular structure and physiological function of chloride channels. Physiol Rev 82: 503-568.

Kasper D, Planells-Cases R, Fuhrmann JC, Scheel O, Zeitz O, et al. 2005. Loss of the chloride channel ClC-7 leads to lysosomal storage disease and neurodegeneration. EMBO J 24: 1079-1091. Kawasaki M, Suzuki M, Uchida S, Sasaki S, Marumo F. 1995. Stable and functional expression of the CIC-3 chloride channel in somatic cell lines. Neuron 14: 1285-1291. Kawasaki M, Uchida S, Monkawa T, Miyawaki A, Mikoshiba K, et al. 1994. Cloning and expression of a protein kinase C-regulated chloride channel abundantly expressed in rat brain neuronal cells. Neuron 12: 597-604. Kida Y, Uchida S, Miyazaki H, Sasaki S, Marumo F. 2001. Localization of mouse CLC-6 and CLC-7 mRNA and their functional complementation of yeast CLC gene mutant. Histochem Cell Biol 115: 189-194.

Neural roles of CLC chloride channels Koch MC, Steinmeyer K, Lorenz C, Ricker K, Wolf F, et al. 1992. The skeletal muscle chloride channel in dominant and recessive human myotonia. Science 257: 797-800. Kornak U, Kasper D, Bosl MR, Kaiser E, Schweizer M, et al. 2001. Loss of the ClC-7 chloride channel leads to osteopetrosis in mice and man. Cell 104: 205-215. Lange PF, Wartosch L, Jentsch TJ, Fuhrmann JC. 2006. ClC-7 requires Ostm1 as a beta-subunit to support bone resorption and lysosomal function. Nature 440: 220-223. Lloyd SE, Pearce SH, Fisher SE, Steinmeyer K, Schwappach B, et al. 1996. A common molecular basis for three inherited kidney stone diseases. Nature 379: 445-449. Poet M, Kornak U, Schweizer M, Zdebik AA, Scheel O, et al. 2006. Lysosomal storage disease upon disruption of the neuronal chloride transport protein ClC-6. Proc Natl Acad Sci USA 103: 13854-13859. Pusch M. 2002. Myotonia caused by mutations in the muscle chloride channel gene CLCN1. Hum Mutat 19: 423-434. Sakamoto H, Kawasaki M, Uchida S, Sasaki S, Marumo F. 1996. Identification of a new outwardly rectifying Clchannel that belongs to a subfamily of the ClC Cl- channels. J Biol Chem 271: 10210-10216. Scheel O, Zdebik AA, Lourdel S, Jentsch TJ. 2005. Voltagedependent electrogenic chloride/proton exchange by endosomal CLC proteins. Nature 436: 424-427. Simon DB, Bindra RS, Mansfield TA, Nelson-Williams C, Mendonca E, et al. 1997. Mutations in the chloride channel

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gene, CLCNKB, cause Bartter’s syndrome type III. Nat Genet 17: 171-178. Steinmeyer K, Klocke R, Ortland C, Gronemeier M, Jockusch H, et al. 1991. Inactivation of muscle chloride channel by transposon insertion in myotonic mice. Nature 354: 304-308. Steinmeyer K, Ortland C, Jentsch TJ. 1991. Primary structure and functional expression of a developmentally regulated skeletal muscle chloride channel. Nature 354: 301-304. Stobrawa SM, Breiderhoff T, Takamori S, Engel D, Schweizer M, et al. 2001. Disruption of ClC-3, a chloride channel expressed on synaptic vesicles, leads to a loss of the hippocampus. Neuron 29: 185-196. Suzuki T, Rai T, Hayama A, Sohara E, Suda S, et al. 2006. Intracellular localization of ClC chloride channels and their ability to form hetero-oligomers. J Cell Physiol 206: 792-798. Thiemann A, Grunder S, Pusch M, Jentsch TJ. 1992. A chloride channel widely expressed in epithelial and nonepithelial cells. Nature 356: 57-60. Uchida S. 2000. In vivo role of CLC chloride channels in the kidney. Am J Physiol Renal Physiol 279: F802-808. Uchida S, Sasaki S. 2005. Function of chloride channels in the kidney. Annu Rev Physiol 67: 759-778. Yoshikawa M, Uchida S, Ezaki J, Rai T, Hayama A, et al. 2002. CLC-3 deficiency leads to phenotypes similar to human neuronal ceroid lipofuscinosis. Genes Cells 7: 597-605.

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IP3 Receptor and Ca2+ Signaling

C. Hisatsune . K. Mikoshiba

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 566

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Characters of Ca2+ Release from IP3Rs in Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 567

3 3.1 3.2 3.3 3.4 3.5 3.6 3.7

Regulation of the IP3 Receptor by Various Associated Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 569 Regulation of Subcellular IP3R Dynamics by 4.1N . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 569 Homer Activity dependently Controls the Coupling Status of mGluR–IP3R Signaling . . . . . . . . . . 570 Cytochrome c Accerelates Apoptosis by Increasing the Channel Activity of IP3Rs . . . . . . . . . . . . . . . . 571 Implication of IP3R-mediated Ca2+ Signaling in Huntington Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 571 Control of IP3Rs by Luminal IP3R-Binding Proteins, ERp44 and Chromogranins . . . . . . . . . . . . . . . . 572 ERp44 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 572 Chromogranin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 573

4 The Physiological Role of IP3R1 in the Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 574 4.1 Disturbed Motor Learning and Motor Coordination in the IP3R1 Mutant Mice . . . . . . . . . . . . . . . . 574 4.2 Abnormal Synaptic Plasticity in IP3R1-Deficient Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 574 5

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Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 575

2009 Springer ScienceþBusiness Media, LLC.

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IP3 receptor and Ca2+ signaling

Abstract: Calcium ions (Ca2+) are ubiquitous second messengers that play an important role in many physiological events including secretion, development, fertilization, and gene expression. However, the proper spatio-temporal regulation of the intracellular Ca2+ concentration is necessary to fulfill the function, and disturbed Ca2+ signaling is known to cause cell death and pathological disease. Inositol 1, 4, 5-trisphosphate receptor (IP3R) is a Ca2+ channel localized on the endoplasmic reticulum (ER) in the many types of cells including neurons, and is a key player to generate the proper intracellular Ca2+ dynamics for cell function. Disruption of IP3Rs leads to various physiological defects including neural development and neural plasticity. Moreover, several lines of evidence indicate that the altered IP3R activity causes supranormal Ca2+ homeostasis leading to various pathological diseases. In this review, we describe how IP3R activity is properly regulated by myriads of associated proteins, and discuss the physiological role of IP3Rs especially in neurons. List of Abbreviations: CTD, C-terminal domain; HAP1A, Htt-associated protein-1A; Htt, Huntingtin; IP3R, Inositol 1, 4, 5-trisphosphate receptor; LFS, low-frequency stimulation; LTD, long-term depression; LTP, long-term potentiation; MSNs, medium spiny neurons; NMDA, N-methyl-D-aspartate; PTP, permeability transient pore

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Introduction

Calcium (Ca2+) is a versatile signal that regulates a myriad of cellular functions, including fertilization, proliferation, development, synaptic plasticity, and gene expression (Berridge et al., 2003). Despite of its importance, the excessive elevation of the intracellular Ca2+ concentration is toxic to cells, and the proper regulations of the intracellular Ca2+ levels in terms of amplitude, space, and duration by various Ca2+ handling proteins (Ca2+ channels, Ca2+ pumps, Ca2+ sequester proteins) are essential for activation of downstream effectors and subsequent successful achievement of the Ca2+-dependent cellular processes. The intracellular Ca2+ level in the resting cells is kept very low (107 M) in contrast to the extracellular space of the cells (2 mM). Stimulation of cells with many extracellular stimuli (e.g., growth factor, neurotransmitter, hormone) increases intracellular Ca2+ levels by two major Ca2+ sources, Ca2+ influx from the extracellular space and Ca2+ release from the internal Ca2+ stores. Inositol 1, 4, 5-trisphosphate receptor (IP3R) is an IP3-gated Ca2+ channels responsible for the Ca2+ release from the internal stores (Furuichi et al., 1989; Berridge, 1993). IP3R is largely composed of the three domains, the NH2-terminal ligand binding domain, the coupling domain, and the COOH-terminal channel domain (Patterson et al., 2004a). Based on the functions on the channel gating, the N-terminal region can be further divided into the N-terminal coupling domain and IP3 binding domain, and the C-terminal channel domain is divided into two region, transmembrane domain and gatekeeper domain (Uchida et al., 2003). There are three distinct subtypes of IP3Rs in mammals (Mignery et al., 1990; Sudhof et al., 1991; Maranto, 1994; Newton et al., 1994; Taylor et al., 1999; Iwai et al., 2005). They show high homology in amino acid sequence (60–80%), but have distinct properties in terms of IP3 affinity, Ca2+ sensitivity, and the regulation by ATP, phosphorylation, and associated proteins (Foskett et al., 2007). To date, a large number of proteins have been shown to directly bind to IP3Rs and regulate the channel activity (Patterson et al., 2004a; Foskett et al., 2007). Most proteins regulate the IP3Rs by interacting with the large cytosolic regions at the NH2- and the COOH-terminus, whereas ERp44 and chromogranins are known to control the channel activity by interacting with the luminal region of IP3Rs (Yoo, 2000; Higo et al., 2005). Moreover, the proportions of these three types of IP3Rs expressed in cells are different and specific to the tissues, e.g., IP3R1 is predominantly expressed in the brain, whereas IP3R2 and IP3R3 are dominantly expressed in pancreas and salivary glands (Taylor et al., 1999; Futatsugi et al., 2005). Thus, the diversity of the IP3R subtype expression, their subcellular localization, and the regulation by IP3, Ca2+, and interacting proteins, all contributes to the complex spatio-temporally coordinated intracellular Ca2+ signaling. In neurons, the Ca2+ release have not been extensively studied compared to the Ca2+ influx, since neurons communicate each other by fast synaptic transmission evoked by action potentials and synaptic deporalization. For example, a large Ca2+ influx through voltage-dependent Ca2+ channels triggers

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neurotransmitter release at synapses. At the postsynaptic sites, N-methyl-D-aspartate (NMDA) receptor is responsible for Ca2+ influx, which is essential for several forms of synaptic plasticity. In addition, synaptic activation generates action potentials which can backpropagate and synchronously induce transient Ca2+ increase at all over the dendrites by opening voltage-dependent Ca2+ channels (Sabatini et al., 2002). However, besides Ca2+ influx mediated by these mechanisms, recent growing evidences indicate that Ca2+ release from the internal Ca2+ store is another important Ca2+ source for generating the neuronal Ca2+ signaling with different spatial and temporal patterns (Barbara, 2002; Ross et al., 2005). There are several interesting features in the neuronal Ca2+ release. First, unlike the transient and small Ca2+ increase through voltage-dependent Ca2+ channels or ligand-gated channels, Ca2+ release has a large amplitude with longer duration, and propagates on the restricted dendrite like ‘‘wave,’’ allowing information to spread long distance, e.g., even into the nucleus. The regenerative property of Ca2+ release relies on the Ca2+ sensitivity of IP3Rs, establishing the Ca2+ wave. In addition, the molecular property of IP3R that it requires both IP3 and Ca2+ for the efficient activation is attractive for neuronal signaling: IP3Rs work as a coincident detector for Ca2+ and IP3 (Iino, 1990; Bezprozvanny et al., 1991; Finch et al., 1991), which may be important for synaptic plasticity. Together with the Ca2+ influx through voltage-dependent Ca2+ channels and ligandgated ion channels, these particular characters of IP3-induced Ca2+ release from the Ca2+ stores can add the complexities to the spatial and temporal patterns of Ca2+ signaling, and regulate a variety of neuronal functions, including synaptic transmission, synaptic plasticity, and gene expression. It is also beginning to be apparent that Ca2+ dyshomeostasis caused by altered IP3R activities are associated with apoptosis and several pathological diseases, including Alzeheimer diseases and Huntington diseases (HDs). These abnormal IP3R activities are likely to be triggered by the enhanced or decreased regulation by associated proteins with mutations. Thus, IP3Rs exist in a macro signal complex with many associated-molecules, whose proper regulation may be important for establishing the coordinated Ca2+ signaling for diverse neuronal functions.

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Characters of Ca2+ Release from IP3Rs in Neurons

Ca2+ release from the ER in response to agonists for the G-protein coupled receptor, e.g., mGluRs and muscarinic receptors, have been reported in the hippocampal neurons from early 1990s (Alford et al., 1993; Jaffe and Brown, 1994; Shirasaki et al., 1994; Seymour-Laurent and Barish, 1995; Miller et al., 1996). Interestingly, IP3R-dependent Ca2+ increase in hippocampal neurons has a distinctive feature: it propagates along the dendrite like ‘‘wave.’’ Application of relatively high concentration of the agonist for mGluR to the apical or basal dendrites triggers local Ca2 transient in the dendrites and the Ca2+ increase subsequently propagates distally and to the soma (Jaffe and Brown, 1994). Similarly, Ca2+ waves are also reported in the neurite of differentiated PC12 cells (Lorenzon et al., 1995). Bath application of bradykinin first focally increases the Ca2+ level at the trigger zone of neurites and then the Ca2+ propagates bidirectionally, to the soma and more apical region. On the other hand, ATP stimulation first increases Ca2+ at the soma of the differentiated PC12 cells, which propagates distally to the apical neurite, suggesting that Ca2+ trigger zones depend on the specific localization of surface receptor. Deporalization by KCl simultaneously increases Ca2+ in the all regions of the neurites, but never elicits Ca2+ wave, indicating the Ca2+ wave do not depend on passive Ca2+ propagation but on regenerative process (Lorenzon et al., 1995). Recently, the mechanism of Ca2+ wave induction in hippocampal neurons was further investigated in more physiological condition. Nakamura et al. found that in the presence of low concentration of mGluR agonist that did not elicit Ca2+ transient by alone, application of action potentials backpropagating in the dendrites induce large Ca2+ waves (Nakamura et al., 1999). Important different characters between the Ca2+ wave and spike-evoked Ca2+ increase are its amplitude and spatio-temporal pattern. The spike-evoked Ca2+ increase occurs at all parts of hippocampal neurons at the same time and is transient, whereas Ca2+ waves triggered by coapplication of spike-mediated deporalization and mGluR activation start from a small part of the proximal apical dendrites and propagate in the dendrites and to the soma (> Figure 31-1) (Nakamura et al., 2000). The peak amplitude of the Ca2+ wave (5  10 mM) is much larger than the linear sum of mGluR-induced Ca2+ release and the spike-evoked Ca2+ transient ( Figure 31-2). The disturbance of these proper functional interactions causes abnormal intracellular Ca2+ homeostasis, leading to pathological diseases and apoptosis. Furthermore, a binding protein, 4.1N, is reported to control the subcellular localization and dynamics of IP3Rs. In the following section, we introduce the molecular mechanism of the regulation of IP3R channels by the recently identified binding proteins, which may be important for the cellular and neural function.

3.1 Regulation of Subcellular IP3R Dynamics by 4.1N Protein 4.1 is a cytoskeleton protein composed of the three domains, the N-terminal membrane binding domain (MBD), the spectrin-actin binding domain (STD), and the C-terminal domain (CTD). Using a yeast two-hybrid screening, two groups independently identified protein 4.1N, a neuronal homolog of the 4.1 family proteins as an IP3R1 binding protein. Zhang et al. found that the last 14 amino acid residues of the COOH-terminus of IP3R1 and the CTD domain of 4.1N are responsible for the binding (Zhang et al., 2003). In addition, they showed that protein 4.1N translocates IP3R1 to the actin filaments at the basolateral membrane domain in polarized Madin–Darby canine kidney cells. Thus, 4.1N plays a critical role in

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. Figure 31-2 Structure of IP3R and its associated proteins. IP3R has five domains: suppressor domain, IP3 binding core domain, coupling domain, transmembrane domain, and coupling domain. Many proteins interact with IP3R and regulate the channel activity of IP3Rs. Black lines show the putative binding region of the associated proteins. IRBIT (Ando et al., 2003), CaM (Yamada et al., 1995; Sienaert et al., 2002), Na+/K+-ATPase (Zhang et al., 2006), FKBP12 (Cameron et al., 1997), Rack1(Patterson et al., 2004b), TRPC3, GAPDH (Patterson et al., 2005), AKAP9 (Tu et al., 2004), PP1 (Tang et al., 2003b)

determining the subcellular distribution of IP3R1. The importance of IP3R1–4.1N interaction in intracellular IP3R1 dynamics was also demonstrated in the hippocampal neurons (Maximov et al., 2003). Fukatsu et al. further visualized the lateral diffusion of GFP-tagged IP3R1 on the ER in the cultured hippocampal neurons and found that the diffusion constant of GFP-IP3R1 depends on the states of actin filaments: disruption of actin filaments increases the diffusion constant and vise versa (Fukatsu et al., 2004). The actin filament-dependent diffusion of GFP-IP3R1 was due to the presence of the 4.1N binding site at the COOH-terminus of IP3R1, and deletion of it from GFP-IP3R1 or coexpression of the CTD domain of 4.1N with GFP-IP3R1 in hippocampal neurons completely abolished the actin dependence of IP3R1 diffusion. The diffusion of neither GFP-IP3R3 which lacks the 4.1N binding site nor GFP-SERCA2a is dependent on actin filaments. Understanding of the physiological role of this regulation of IP3R1 diffusion in neuronal function, e.g., synaptic plasticity, will be expected.

3.2 Homer Activity dependently Controls the Coupling Status of mGluR–IP3R Signaling Homer family proteins are scaffolding proteins, and composed of the three subtypes of Homer 1, 2, and 3 in mammal (Xiao et al., 2000). They contain two major domains, a highly conserved EVH1 domain and a coiled-coil domain. Whereas the coiled-coil domain contributes to the homo- and hetero-tetramer formation (Hayashi et al., 2006), the EVH1 domain recognizes and interacts with a consensus homerbinding motif ‘‘PPXXF’’ of various target proteins including IP3R1 (Tu et al., 1998) and mGluR1a (Brakeman et al., 1997). Consequently, Homer can physically crosslink IP3R1 with mGluR1a, forming a

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large signal complex. The IP3R1–Homer–mGluR complex facilitates the Ca2+ signaling in response to the mGluR stimulation in Purkinje cells, and disruption of the complex affects Ca2+ signaling (Tu et al., 1998). Interestingly, the formation of the multimeric complex can be regulated by synaptic activity through the control of the expression level of Homer 1a, a splicing variant form of Homer 1. Because Homer 1a lacks the coiled-coil domain (Kato et al., 1997; Xiao et al., 1998), Homer 1a could neither form tetramer nor make a physical link between homer-associated proteins, working as a dominant negative form of Homer proteins (Xiao et al., 2000). The physiological effects of the regulation of the multimeric complex by Homer 1a on synaptic plasticity and higher brain function is expected to be clear in the future experiments.

3.3 Cytochrome c Accerelates Apoptosis by Increasing the Channel Activity of IP3Rs The activity of IP3R1 is biphasically regulated by cytosolic calcium levels: the activity of IP3R1 increases in a Ca2+-concentration-dependent manner, but decreases when the cytosolic Ca2+ concentration beyond 300 nM (Bezprozvanny et al., 1991). This bell-shaped activity of IP3R1 in response to the intracellular Ca2+ level is probably one of the important feed back mechanisms that protect the cells from the excessive Ca2+ release through IP3Rs. By yeast two-hybrid screening, Boehning et al. identified cytochrome c as a binding protein of the cytosolic COOH-terminus of IP3R1 (Boehning et al., 2003). They found that cytochrome c released from mitochondria through permeability transient pore (PTP) translocates to the ER, and binds IP3R1 early in apoptosis. Importantly, cytochrome c binding diminishes the bell-shaped response of IP3R1 to Ca2+ and leaves IP3R1 open even in the higher cytosolic Ca2+ concentration. The resultant activation of IP3R1 leads to further mitochondrial Ca2+ overload and cytochrome c release, resulting in the amplification of the apoptotic signals (Boehning et al., 2003). Thus, the positive regulation of IP3R by cytochrome c is likely a feed-forward mechanism during early apoptosis. More recently, the critical role of the interaction of cytochrome c to IP3R1 is also shown in more physiologic apoptotic cell death using a cell permeable inhibitor peptide (Boehning et al., 2005). In the study, introduction of the 16-amino acid peptide, encoding a cluster of glutamic acid residues of the C-terminus of IP3R1 in Jurkat cells, which inhibits the IP3R1–cytochrome c interaction was shown to be enough for inhibition of Fas ligand-induced apoptotic cell death. In future, the cell permeable peptide would be expected to be a good therapeutic tool for many diseases associated with apoptotic cell death, such as inflammatory disturbances and neurodegenerative diseases.

3.4 Implication of IP3R-mediated Ca2+ Signaling in Huntington Disease HD is an autosomal-dominant neurodegenerative disorder with chorea and psychiatric disturbance. The notable feature of HD is a selective and progressive loss of medium spiny neurons (MSNs) in the caudate and putamen (Vonsattel et al., 1985; Vonsattel and DiFiglia, 1998). HD is caused by abnormal polyglutamine (polyQ) expansion (>35) of the N-terminus region of Huntingtin (Htt) (The Huntington’s Disease Collaborative Research Group, 1993). Deletion of Htt in mice leads to embryonic lethal, suggesting a critical role of Htt in development (Duyao et al., 1995; Nasir et al., 1995); however, precise role of Htt is not known. Although Htt is ubiquitously expressed in the brain and also in nonneuronal tissues, the loss of cells caused by polyQ expanded Htt (Httexp) are selectively observed in MSNs, and the reason remains unknown. There have been many studies suggesting the toxic function of Httexp, in the cellular events such as gene transcription, induction of apoptosis, and synaptic transmission (Tobin and Signer, 2000; Rubinsztein, 2002; Sugars and Rubinsztein, 2003; Li and Li, 2004). In addition to these biological defects by Httexp, several lines of evidence also indicate the deregulated intracellular Ca2+ homeostasis caused by Httexp (Bezprozvanny and Hayden, 2004). Although there are several potential mechanisms that lead to disturbed intracellular Ca2+ homeostasis in HD such as abnormal mitochondrial Ca2+ handling

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(Panov et al., 2002) and potentiation of NMDA receptor activity by Httexp (Chen et al., 1999; Zeron et al., 2001), we described here the potential involvement of the enhanced IP3R1 activity by Httexp. Using a yeast two-hybrid screening system, Tang et al. identified Htt-associated protein-1A (HAP1A) as an IP3R1 binding protein (Tang et al., 2003a). HAP1A associates with the COOH-terminal cytosolic region of IP3R1. In addition, ternary complex of IP3R1–HAP1A–Htt was demonstrated both in vitro and in the neurons within rat brain. HAP1A promotes association of Htt to IP3R1 and facilitates the channel activity of IP3R reconstituted in planer lipid bilayers. Furthermore, disease-related Httexp but not normal Htt by itself increases IP3R1 channel activity by sensitizing IP3R1 to activation by submaxial doses of IP3 even without HAP1A. They further showed the functional effect of Httexp on Ca2+ homeostasis in MSNs. Rat MSNs expressing disease-associated Httexp showed relatively higher resting Ca2+ level and the enhanced metabotropic glutamate receptor 1 and 5 (mGluR1/5)-driven Ca2+ signals (Tang et al., 2003a). More recently, a role of disturbed IP3R-mediated Ca2+ signaling during apoptosis in HD was directly demonstrated; the inhibitors of IP3Rs inhibit apoptosis of MSNs from transgenic mice expressing diseaseassociated Httexp (Tang et al., 2005). Mitochondria are known to closely communicate with the ER and actively accumulate Ca2+ released from IP3Rs upon physiological and pathophysiologic condition (Rizzuto et al., 1998; Csordas et al., 1999). In addition to the caspase and calpain activation by supranormal intracellular Ca2+ level of MSNs by NMDARs and IP3Rs, increased Ca2+ release through IP3R1 actviated by Httexp could lead to abnormal Ca2+ uptake into mitochondria, leading to the activation of mitochondrial PTP, cytochrome c release, activation of calpain and caspase, and ultimately induction of apoptosis.

3.5 Control of IP3Rs by Luminal IP3R-Binding Proteins, ERp44 and Chromogranins IP3R has six membrane-spanning regions in the channel domain, being three loops residing in the ER lumen. Based on the primary sequences, the largest third loop is further divided into two subdomains, first half region called ‘‘L3V domain’’ and latter one ‘‘L3C domain’’ containing the channel pore. The L3C domain is highly conserved among the three types of IP3Rs. In contrast, the L3V region is highly divergent and thus expected to have a specific meaning, e.g., a subtype-specific regulation by interacting protein. Chromogranin and ERp44 are ER-resident proteins and have recently shown to bind to the L3C and L3V regions of IP3R, respectively. These proteins sense intraluminal environment of the ER, especially Ca2+ concentration, and can control the Ca2+ concentration of ER by regulating the IP3R channel activity. Since these proteins are highly expressed in the brain and disruption of ER Ca2+ homeostasis leads to ER stress with abnormal posttranslational protein folding and modification, which regulates neuronal survival, the regulation of IP3Rs by these molecules may be involved in the pathogenesis of various neurodegenerative diseases.

3.6 ERp44 ERp44 is an ER luminal protein that belongs to a thioredoxin (TRX) family protein (Anelli et al., 2002). Using an affinity column using L3V region of IP3R1 as a bait, Higo et al. identified ERp44 as an IP3R1 binding protein. Interestingly, ERp44 binding to IP3R1 is dependent on PH, redox state, and Ca2+ concentration (Higo et al., 2005). ERp44 binds IP3R1 at pH = 5.2, but not at pH = 7.5. However, ERp44 can interact with IP3R1 even at pH = 7.5, if the Cys residues of the L3V region of IP3R1 are reduced. The association of IP3R1–ERp44 is diminished if Ca2+ concentration is increased. In contrast to the enhancement of IP3R activity by chromogranin association (see later), the effect of ERp44 is inhibition of the channel activity of IP3R1. As expected from the divergent L3V sequences among three types of IP3Rs, ERp44 selectively binds to and inhibit IP3R1, but not IP3R2 and IP3R3. This is supported by the

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finding that ERp44 inhibited the Ca2+ release and the subsequent apoptosis in chicken DT40-KMN60 cells expressing only IP3R1, but not in the other mutant DT40 cells lacking IP3R1 (DT40-1KO) upon BCR stimulation (Higo et al., 2005). Since depletion of Ca2+ store triggers caspase activation in some Ca2+dependent apoptosis (Nakagawa and Yuan, 2000; Nakagawa et al., 2000), ERp44 may keep the ER store not depleted and avoid ER-dependent apoptosis by inhibiting the IP3R1 activity in the situation in which Ca2+ contents of the ER is getting decreased.

3.7 Chromogranin Chromogranin A and B are high capacity, low affinity Ca2+ storage proteins (CGA: 30–50 mol of Ca2+/mol, Kd = 4 mM at pH = 5.5; CGB: 100 mol of Ca2+/mol, Kd = 1.5 mM at pH = 5.5) that are highly present in the secretory granules but also in the ER of endocrine cells, neurons, and neuroendocrine cells (Yoo and Albanesi, 1991; Yoo et al., 2001). Because of the high Ca2+ capacity and high concentration (1  2 mM) of chromogranins in the secretory granules, the secretory granules have been thought to function as Ca2+ stores in endocrine cells (Yoo, 2000). This is directly demonstrated by the findings that the isolated secretory granules from adrenal medullary chromaffin cells (Yoo and Albanesi, 1990) and pancreatic acinar cells (Gerasimenko et al., 1996) were able to release Ca2+ in response to IP3. In addition, Ca2+ release from the secretory granules in the goblet cells has been shown to regulate the cytoplasmic Ca2+ concentration (Nguyen et al., 1998), which is important for the process of exocytosis (Maruyama et al., 1993; Maruyama and Petersen, 1994; Mundorf et al., 1999; Mundorf et al., 2000). Therefore, understanding the regulatory mechanism by which IP3-sensitive Ca2+ release occurs from the secretory granules is an important issue. Several studies have shown that chromogranin directly interacts with and regulates IP3Rs in a pH-dependent manner (Yoo et al., 2000). CGA binds to the L3C region of IP3Rs at pH = 5.5, which is almost similar to the intragranular pH of the mature secretory granules, but not at pH = 7.5 (Yoo and Lewis, 1998). The CGA binding to IP3R was shown to increase the channel open probability by approximately tenfold without the change of the amplitude at the intraluminal pH = 5.5. But the effect of CGA was abolished when intraluminal pH was changed to 7.5, the condition where no interaction of CGA to IP3R occurred (Thrower et al., 2002). In contrast to CGA, CGB more strongly binds to IP3R at pH = 5.5 and also even at pH = 7.5 albeit in a lower affinity compared with the binding at pH = 5.5 (Yoo et al., 2000). The CGB–IP3R interaction markedly increases the mean open times and open probabilities (20-fold) of IP3R channel activity at the intraluminal pH = 5.5. As a reflection of the CGB binding to IP3R at pH = 7.5, the effect of CGB on the IP3R channel activity was still observed at the luminal pH = 7.5 (Thrower et al., 2003). The CGB–IP3R interaction and its functional effect on IP3R channel activity at a physiological pH imply the regulation of IP3R by CGB in the ER, where pH is considered to be almost neural and stable. Consistent with the idea, Choe et al. reported that the expression of an N-terminal 20-amino acid fragment of CGB that can disrupt the interaction between CGB with IP3R results in disturbed intracellular Ca2+ signaling triggered by carbachol in differentiated PC12 cells (Choe et al., 2004). Furthermore, Jacob et al. reported a potential role of chromogranin in the establishment of the Ca2+ trigger zone in the proximal dendrites of PC12 cells (Jacob et al., 2005). They analyzed the subcellular distribution of proteins related to the IP3-mediated Ca2+ signaling in PC12 cells and found that chromogranin shows specialized distributions, highly accumulation in the neurites. Whereas Ca2+ wave initiates from ‘‘trigger zone’’ on the dendrites and propagates to the soma in the control peptide expressing PC12 cells, the expression of 20-amino acid CGB fragment reversed the order: Ca2+ increases first in the soma which follows the Ca2+ increase in the neurites. Intriguingly, the abnormal expression level of chromogranin has been reported in the brains with several human diseases, such as schizophrenia, Alzheimer’s disease (Nishimura et al., 1994; Yasuhara et al., 1994; Marksteiner et al., 2000). Although we do not know any relationship between the abnormal expression of chromogranin with the causes of these diseases at present, disturbed IP3-induced Ca2+ signaling caused by changes of chromogranin expression levels might be one of the causes of these diseases and related to the disease progression.

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The Physiological Role of IP3R1 in the Brain

Quite a lot of information about the physiological roles of IP3R1 in the central nervous system has been revealed by the analysis of the IP3R1 knockout mice and spontaneous opisthotonos (opt) mutant mice, which demonstrates the critical role of IP3R1 in motor learning and motor coordination.

4.1 Disturbed Motor Learning and Motor Coordination in the IP3R1 Mutant Mice Most of the mutant mice with disruption of ip3r1 gene die in utero by unknown reasons, suggesting the critical role of IP3R1 in development (Matsumoto et al., 1996). Successfully born mutant mice however appear normal, but show retarded growth. They begin to show ataxia with loss of balance while standing and walking around postnatal day 9 (P9), and exhibit truncal torsions by P15. Thereafter, they display tonic or tonic-chronic seizures and die by the weaning period. Despite of such a severe phenotype, the gross anatomy of the mutant brain is not significantly different from normal mice, although the abnormality of dendritic morphology of Purkinje cells with reduced branches and synaptic vesicle accumulation at PF–Purkinje cell synapse were recently found in the mutant mice (Hisatsune et al., 2006). Purkinje cells from the IP3R1-deficient mice lacked mGluR-mediated Ca2+ release and showed impaired LTD on the PF–Purkinje cell synapse in cerebellum (Inoue et al., 1998). The spontaneous opt mutant mice also display epileptic-like behavior and die within 4 weeks (Street et al., 1997). Genomic and molecular analysis revealed the alternation of IP3R1 gene in the opt mutant mice. In the mutant mice, the deletion of two exons of IP3R1 gene occurred without interruption of the translational reading frame. Consequently, the opt mutant mice expressed the truncated form of IP3R1 at markedly reduced levels, which lacks a part of the regulatory domain containing phosphorylation sites by protein kinase A (PKA) and protein kinase G (PKG) and potential ATP binding sites. Interestingly, despite of the reduced IP3R1 protein level, Purkinje cells from the opt mutant mice elicit a comparable Ca2+ release from the intracellular stores compared with control Purkinje cells, although repeated agonist stimulation result in less attenuation of Ca2+ release (Street et al., 1997). Apparently, restrict regulation of IP3R activity, e.g., by phosphorylation and ATP, seems to be necessary to proper Ca2+ signaling in Purkinje cells and higher brain function.

4.2 Abnormal Synaptic Plasticity in IP3R1-Deficient Mice LTD, a synaptic plasticity at the PF–Purkinje cell synapse in cerebellum is an important cellular basis of motor learning and motor coordination. In the cerebellum of the IP3R1 knockout mice, LTD is completely abolished (Inoue et al., 1998). Purkinje cells from the IP3R1 mutant mice show no Ca2+ release from the ER, but normally depolarization evoked Ca2+ transient. In addition, introduction of functional blocker ‘‘antibody for IP3R1’’ also inhibit LTD induction. Thus, this study provide the conclusive evidence that Ca2+ release from IP3R1 is an essential event in the induction of LTD, and the defect of induction of cerebellar LTD would explain the disturbed motor learning and coordination observed in IP3R1 knockout mice. In support with this observation, the mice lacking metabotropic glutamate receptor 1 (mGluR1) and Gqa (Kleppisch et al., 2001; Hartmann et al., 2004), both of which activation is known to lead the IP3 production and subsequent IP3R-induced Ca2+ release, showed a similar motor deficit and the defect of LTD induction (Aiba et al., 1994). IP3R1 also contribute to the synaptic plasticity in the hippocampus neurons, which is profoundly associated with learning and memory. Several types of synaptic plasticity have been known in hippocampus, such as long-term potentiation (LTP), LTD, depotentiation (DP) by which low-frequency stimulation (LFS) reverses a pre-established LTP, and LTP suppression, a phenomenon that application of LFS prior to the LTP induction abolishes LTP. In the CA1 hippocampal neurons of IP3R1 knockout mice, facilitation of

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LTP induced by short tetanus (10 pulses at 100 Hz) is observed at CA1 synapses, whereas basal synaptic transmission is normal compared with control mice. In addition, DP and LTP suppression are also abolished in the CA1 hippocampal neurons of IP3R1 knockout mice (Fujii et al., 2000). Interestingly, facilitation of LTP and DP abolishment observed at the CA1 synapse in IP3R1 knockout mice are dependent on the activity of NMDARs. Nishiyama et al. further demonstrates the contribution of IP3R1 in the input specificity of the activity-dependent synaptic plasticity: deletion of IP3R1 results in the complete lack the heterosynaptic LTD in the CA1 region of hippocampus (Nishiyama et al., 2000). It is an interesting issue to investigate whether Ca2+ wave mediated by IP3R1 in hippocampal neurons contributes to the input specificity of the activity-dependent synaptic plasticity.

5

Concluding Remarks

During the last decades, there is a great advance in identifying the molecular players that regulate IP3Rs, indicating the presence of a macro signal complex of IP3Rs. Because the expression patterns of those regulatory proteins were different from one tissue to others, the members constituting the macro signal complexes should be divergent, and the effect of the interaction of an IP3R-binding protein on IP3Rs may be distinct in a tissue-specific manner. Therefore, it will be necessary to analyze the molecular interplay of all these binding proteins to understand their effect on IP3Rs in vivo. In addition, it may be also possible that IP3R affects the function and localization of many proteins constituting the macro signal complex in an independent fashion of its channel activity. Disruption of IP3R1 gene reveals the critical roles of IP3R1 in the brain function. However, the mechanism by which IP3R1 regulates the brain function is largely unknown. It is an important future issue to understand the relationship between the IP3R-mediated Ca2+ signaling in a specific part of the brain and the brain function (learning, memory, emotion, and consciousness, disease such as epilepsy and schizophrenia).

Acknowledgement We thank Dr. Akihiro Mizutani and Dr. Yukiko Kuroda for fruitful comments on the manuscript. Supported by grants from the Ministry of Education, Science, and Culture of Japan (K. M. and Y. N.), Grant-in-Aid for Young Scientists (C. H.), and the Japan Science and Technology Agency.

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E. Carafoli . D. Lim

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Historical Notes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 582

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General Properties of the PMCA Pump . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 582

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Primary Structure of the PMCA Pump . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 583

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Membrane Topology and Functional Domains of the PMCA Pump . . . . . . . . . . . . . . . . . . . . . . . . . . . 583

5 Isoforms of the PMCA Pump . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 584 5.1 Distribution of PMCA Isoforms in the Adult Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 584 5.2 Developmental Changes in the PMCA Distribution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 585 6 Splicing Variants of the PMCA Pumps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 586 6.1 Functional Diversity of the PMCAs: Structure–Function Relationships . . . . . . . . . . . . . . . . . . . . . . . . . . 587 7

Regulation of the Activity of the PMCA Interaction with Modulators . . . . . . . . . . . . . . . . . . . . . . . . . 588

8 8.1 8.2 8.3

PMCA Pumps and Pathology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 590 Studies on PMCA Knock-Out Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 590 PMCA2 Mutants and Hereditary Hearing Loss . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 590 PMCA Pumps in Other Pathological Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 591

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Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 592

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Abstract: To function properly, cells must maintain a very low cytosolic free Ca2+ concentration (50–200 nM) in the face of an extracellular concentration, which is 1–2 mM, and can reach even 10 mM in sea water, which is the extracellular ambient of some cells, e.g., oocytes, of some marine species. To maintain such a steep concentration gradient, eukaryotic cells have developed efficient systems for Ca2+ extrusion. Two of them operate in the plasma membrane, a Ca2+ ATPase (PMCA pump), and a Na+/Ca2+ exchanger (NCX), two inside the cell, in the endo(sarco)plasmic reticulum (ER) (a Ca2+ ATPase, the SERCA pump), and in the secretory pathway (the SPCA pump). The Ca2+-mediated signal transduction requires the concerted operation of all these systems, to eventually offset the penetration of Ca2+, which occurs through a number of specific channels. The final result of the operation is the precise tuning of internal Ca2+ and the control of its elevation in time and space to activate (or terminate) most life processes, beginning with the origin of new cell life at fertilization. When the operation of the Ca2+ controlling systems becomes unbalanced and fails to clear abnormal cytosolic Ca2+ concentrations (cell Ca2+ overload), irreversible activation of the death pathways occurs, condemning cells to death. Of the two plasma membrane exporting systems one (the NCX) functions with low Ca2+ affinity, but high Ca2+ exporting capacity, the other (the PMCA) with opposite properties, i.e., with low Ca2+ transporting capacity, but very high Ca2+ affinity. The PMCA pump can thus be considered as the fine tuner of cytosolic [Ca2+]. This chapter describes its properties, paying particular attention to aspects where important developments have recently occurred, e.g., regulation, and structure–function relationships.

1

Historical Notes

Forty years have elapsed since the discovery by Schatzmann of the ATP-dependent Ca2+ ejection from red blood cells (Schatzmann, 1966). More than a decade after the discovery of the process the protein responsible for it (the PMCA pump) was isolated using a calmodulin affinity column (Niggli et al., 1979). After another decade the pump was eventually cloned and the deduced amino acid sequence showed that it repeated the membrane topology of the other pumps of this type, which were already known at that time (Shull and Greeb, 1988; Verma et al., 1988). The purification and the cloning work heightened interest on the pump and were followed by numerous studies that established its properties, its regulatory aspects and, in recent times, the consequences of its dysregulation.

2

General Properties of the PMCA Pump

The PMCA is a P-type pump, i.e., it is characterized by the formation of a high-energy aspartyl-phosphate intermediate that is coupled to the translocation of the Ca2+ and to the transition of the enzyme between E1 and E2 conformational states. In the E1 state the enzyme binds Ca2+ with high affinity to a Ca2+-binding site presumably accessible to the cytosolic side. The phosphorylation of the catalytic aspartate is coupled to the E1P -> E2P transition, which exposes the bound Ca2+ to the extracellular face and decreases the affinity of the binding site, promoting the release of Ca2+. This occurs before the cleavage of the E2P intermediate. After that, the enzyme returns to the E1 conformation. To achieve high affinity for Ca2+ the PMCA must be activated, since the nonactivated pump has low 2+ Ca affinity (Km > 10 mM). The Ca2+ affinity of the pump is increased to a Km of about 0.2 mM by the exposure to a number of regulators, the most important probably being calmodulin (Carafoli, 1992) and acidic phospholipids (Niggli et al., 1981). Phosphorylation reactions by PKA and PKC kinases (Caroni and Carafoli, 1981; Wang et al., 1991) also increase Ca2+ affinity, as does the self-association of pump units (Kosk-Kosicka and Bzdega, 1988), and the C-terminal cleavage by calpain (James et al., 1989). More detail on the regulation of the PMCA pump can be found below.

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As all P-type pumps, the PMCA is inhibited by the phosphate analog orthovanadate (VO3(OH)2
) and by La3+. The inhibition by La3+ has an interesting peculiarity with respect to other P-type ATPases, i.e., La3+ increases the steady-state level of the phosphorylated intermediate blocking the E1P -> E2P transition (Quist and Roufogalis, 1975; Szasz et al., 1978; Luterbacher and Schatzmann, 1983). In all other P-type pumps La3+reduces it. This allows the identification of the phosphorylated PMCA pump in preparations containing much higher amounts of other P-type pumps, e.g., the SERCA pump. This feature is particularly useful, since the amount of the PMCA protein in most eukaryotic cells is very limited, i.e., it does not exceed 0.1% of the total membrane protein, whereas other P-type pumps may be much more abundant. At variance with the SERCA pump, the PMCA hydrolyses 1 ATP molecule per 1 Ca2+ ion (Niggli et al., 1982; Hao et al., 1994). It is partially electrogenic and exchanges 1 Ca2+ for 1 H+ (Hao et al., 1994).

3

Primary Structure of the PMCA Pump

The complete amino acid sequence of the PMCA pump was deduced using rat brain (Shull and Greeb, 1988) and human teratoma (Verma et al., 1988) cDNA libraries, respectively. The described sequences of two rat brain isoforms contained 1176 and 1198 amino acids. The two pumps were denominated PMCA1 and PMCA2, respectively. The human sequence shared more than 99% similarity with rat PMCA1 in the first 1117 residues, but was significantly different in the remainder of the sequence. Later on, it was found that the sequence discrepancy was due to a complex alternative splicing process (Strehler et al., 1989), which generated more than 20 splicing variants for the four basic isoforms of the pump that were eventually cloned (Strehler and Zacharias, 2001). It was indeed soon found that the pump is the product of four different genes, ATP2B1 to ATP2B4 (for PMCA1 to PMCA4, respectively), which in humans are located in the following chromosomal loci: 12q21-q23, 3p25-p26, Xq28, and 1q25-q32 (Olson et al., 1991; Brandt et al., 1992a; Latif et al., 1993; Wang et al., 1994). The number and size of coding exons are conserved among the four isoforms, although the sizes of the human genes ranges from 60 kb for PMCA3 to over 350 kb for PMCA2 (Brandt et al., 1992b). To avoid confusion, the notation “isoforms” will be used in this contribution for the products of the four genes, the notation “splice variants” for the variants generated by the alternative splicing events of each one of the four primary transcripts.

4

Membrane Topology and Functional Domains of the PMCA Pump

The analysis of the membrane topology of the PMCA pump as deduced from the primary sequence revealed 10 membrane-spanning domains (TM1 to TM10, > Figure 32‐1) with the N- and C-termini located in the cytosol. The 10 TM domains are connected by five short extracellular loops, about 80% of the mass of the protein thus protruding into the cytosol. The N-terminal cytoplasmic domain, encompassing the first 80–90 amino acids, is the pump portion that varies most among the four basic isoforms. The intracellular loop between TM2 and TM3 is predicted to be mainly composed of b-sheets and contains a stretch of basic amino acids, which is one of the two binding sites for activatory phospholipid (PL) in the pump, the other site being the basic, C-terminal calmodulin-binding domain. The loop also contains one of the two receptor sites for the calmodulin-binding domain (see > Figures 32‐1 and > 32‐3). The large intracellular loop between TM4 and TM5 contains the other receptor site for the calmodulin-binding domain and the catalytic domain of the pump (the invariant Asp, which is Asp495 in isoform PMCA4), and the ATP (FITC)-binding site (see > Figure 32‐1). The long C-terminal tail is the major regulatory domain of the pump, as it contains the calmodulin-binding domain, phosphorylation sites for PKA and PKC, and the binding sites for a number of regulatory proteins (see below).

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. Figure 32‐1 Membrane topology of the PMCA pump. The ten transmembrane domains (TM) are symbolized by cylinders. The autoinhibitory calmodulin-binding domain is bound to receptor sites in the first (between TM2 and TM3) and the second (between TM4 and TM5) intracellular loops of the pump (oval). Splice sites A and C are depicted as grey cylinders. D, the invariant Asp in the catalytic site of the pump; K, the Lys labeled with FITC (ATP) in the ATP-binding site

5

Isoforms of the PMCA Pump

The products of the same PMCA gene share up to 95% sequence homology in different mammalian organisms, but significant differences exist instead among the four gene products in the same species (70–80% homology). Significant differences also exist in tissue distribution. The PMCA1 and PMCA4 pumps are ubiquitously expressed in animal tissues and are thus thought to be housekeeping isoforms. PMCA1 is expressed most abundantly in brain, lung, and intestine (Greeb and Shull, 1989; De Jaegere et al., 1990; Howard et al., 1993; Stauffer et al., 1995), PMCA4 in kidney, erythrocytes, skeletal muscle, heart, stomach, intestine, brain and spermatozoa (Brandt et al., 1992b; Howard et al., 1993; Stauffer et al., 1995; Okunade et al., 2004). The PMCA2 and PMCA3 isoforms have much more restricted tissue distribution, suggesting functional specificity. In humans the PMCA2 pump is essentially a brain isoform (Stauffer et al., 1995), whereas PMCA3 is expressed in brain and skeletal muscles (Greeb and Shull, 1989; Stauffer et al., 1995; Brown et al., 1996). The PMCA2 and PMCA3 isoforms are the most abundant in rat pancreatic b-cells (Kamagate et al., 2000). The splice variants of PMCA2 are most abundant in brain and heart (Greeb and Shull, 1989).

5.1 Distribution of PMCA Isoforms in the Adult Nervous System In the nervous system, the PMCA isoforms exhibit both a regional and a developmental distribution pattern. The transcript for PMCA1 is predominant in the CA1 pyramidal cells of the hippocampus (Stahl et al., 1992) but high levels of it were also found in the forebrain, especially in the paraventricular and supraoptic nuclei, and in the subependymal layer of the diencephalon and CA1 hippocampus, in midbrain

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(oculomotoric nucleus), and in hindbrain, especially in the locus ceruleus and the nucleus subceruleus of the myelencephalon (Stauffer et al., 1997; Burette et al., 2003). In human brain the PMCA1 isoform is mostly present in the cerebellum and the brainstem (Stauffer et al., 1995), but in the hippocampus the predominant isoforms are PMCA2 and PMCA4 (Zacharias et al., 1997). The mRNA for PMCA2 was found in the cortex, cerebellum, Purkinje cells, and brainstem (Stahl et al., 1992; Stauffer et al., 1995). The PMCA2 isoform was confirmed at the protein level in the Purkinje cells, where it appears to be the only isoform expressed in the dendritic spines (Stauffer et al., 1997), and in the outer hair cells of the inner ear (Dumont et al., 2001). The PMCA2 isoform was also detected in the cerebral cortex, in the striatum, and in the olfactory bulb and the hippocampus (Burette et al., 2003). Early in situ hybridization studies of tissue distribution of the PMCA3 isoform had located it to the habenula and the choroid plexus (Stahl et al., 1992), but the isoform was then also found in the cerebellum, in the cortex, and in the hippocampus (Eakin et al., 1995; Stauffer et al., 1997; Burette et al., 2003). High levels of PMCA4 mRNA were found in the piriform cortex, especially in the amygdaloid nucleus and laminae 2 and 6 of the cerebral cortex (Stahl et al., 1994), and in the cerebellum (Stauffer et al., 1995; Guerini et al., 1999). Specific antibodies have confirmed the widespread expression of the PMCA4 isoform in the brain, indicating that at least two splice variants (4a and 4b) are coexpressed in most brain regions (Burette et al., 2003). PMCA4 expression is most abundant in the cerebral cortex, accessory olfactory bulb, anterior olfactory nucleus, bed nucleus of the stria terminalis, supraoptic nucleus, and spinal trigeminal nucleus (Burette et al., 2003). In glial cells of the brain cortex three isoforms, PMCA1, 2, and 4 were found (Fresu et al., 1999), although only PMCA1 and 2 were found in cultured C6 glioma cells.

5.2 Developmental Changes in the PMCA Distribution The majority of the studies concerning the spatial distribution of the PMCA isoforms have been performed on adult tissues. The developmental peculiarities of their distribution in the cerebellum have been studied in recent work by Sepulveda and coworkers (Sepulveda et al., 2005; Sepulveda et al., 2007) in chicken. A comprehensive study of the developmental pattern of PMCA expression by in situ hybridization on sections of mouse embryos has shown that the PMCA1 isoform could be detected ubiquitously in the embryo at day 9.5 (Zacharias and Kappen, 1999). The PMCA1 was ubiquitously expressed during development in spite of small fluctuations in the level of expression. Thus, this isoform can indeed be considered as a “housekeeping” isoform. All other isoforms only appeared around day 12.5 and exhibited marked changes during the successive embryonic development. PMCA2 was confined to the nervous system, in the peripheral nervous system the highest levels being detected in the dorsal root ganglia and the retinoblasts of the developing eye. In the central nervous system the most intense labeling was found in the external granular layer of the cerebellum, which is the site of developing Purkinje neurons, and in the interpeduncular nucleus. The transcript of PMCA3 was widely expressed in tissues between days 12.5 and 15.5, but at later developmental stages the distribution of this isoform became restricted to the nervous system, the developing limb skeletal muscles, and the lung. Strong expression was also found in the choroid plexus, which is almost the only site of PMCA3 expression in adult organisms (see above). The PMCA4 isoform was expressed at lower levels from day 12.5 to day 18.5. Its distribution was ubiquitous and about the same in all tissues, with the exception of liver, where its expression was higher. At later time points the PMCA4 mRNA was most abundant in the brain, dorsal root ganglia, and the intestine, approaching the distribution seen in the adult mouse tissues (Strehler and Zacharias, 2001). Variations of the developmental expression pattern with respect to mice have been detected in rats and chickens (Brandt and Neve, 1992). For example, in rat embryos PMCA2 appeared much later, whereas in chickens PMCA3 appeared at high levels in the cerebellum at early embryonic stages, and remained constant during the development (Sepulveda et al., 2007). PMCA3 appears to be the most abundantly expressed isoform in chick cerebellum (Sepulveda et al., 2007). In relating the developmental aspects of the distribution and expression of the PMCA isoforms to activity, the expression of endogenous activators must also be considered, calmodulin being the most important. Calmodulin expression has been shown to increase in rat brain and other rat tissues in the late embryonic and early postnatal period (Rainteau et al., 1988; Weinman et al., 1991).

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Splicing Variants of the PMCA Pumps

As mentioned earlier, each PMCA gene produces several matured mRNA variants (splice variants) (Shull and Greeb, 1988; Strehler et al., 1989). The variants are generated at two sites named site A and site C (Strehler and Zacharias, 2001). An additional splicing site in the C-terminal portion of the pump (site B) is now considered to be aberrant. Two different nomenclature systems have been proposed to identify the PMCA splice variants (Keeton et al., 1993; Carafoli, 1994) (see > Table 32‐1). The most commonly used

. Table 32‐1 Nomenclature of splice variants of PMCA pumps

Splice site A

Splice site C

Original nomenclature (Keeton et al., 1993) z x y w v a b c d e f

Alernative nomenclature (Carafoli, 1994) AI AII AIV AIII AV CII CI CIII CIV CV CIV

system (Keeton et al., 1993) names the variants produced at site A with letters from z to w according to the number of inserted exons, the shortest, in which no insertions occur, being variant z. The site C variants are designated with letters from a to d, the b variant having no inserts (see > Figure 32‐2). Splice site A is located in the cytoplasmic loop between TM2 and TM3 upstream of the phospholipid binding site, and variants at this site are generated by the insertion of one extra exon of 39 nt in PMCA1, of one of 42 nt in PMCA2 and 3 and of one of 36 nt, which correspond to exon 8 of PMCA3 gene, in PMCA4 (Burk and Shull, 1992), generating splice variant x for all three isoforms (> Figure 32‐2). In PMCA1 the 39 nt exon is apparently never spliced out (Carafoli, 1994). However, the splicing of the PMCA2 transcript at site A is more complex, as it may include two additional exons (33, and 60 nt) corresponding to exons 6 and 7 of the PMCA2 gene. The insertion of both the 33 nt and the 60 nt exons in PMCA2 generates variant y, and the insertion of all the 3 exons generates variant w. Splicing at site A does not induce loss of reading frame, and thus does not modify the remainder of the translated protein. Splicing site C is located in the calmodulinbinding domain in the C-terminal tail of the PMCA and includes the insertion of homologous exons of 154, 172, 154 and 175 nt in the PMCA1 to 4 isoforms, respectively (Strehler et al., 1989; Adamo and Penniston, 1992; Heim et al., 1992; Brandt et al., 1992b; Stauffer et al., 1993). The insertion corresponds to exon 23 of rat PMCA3 (Burk and Shull, 1992). The inserted exons might have internal donor splice sites, generating additional variants, of which 2 were found for PMCA1 (87 nt and 114 nt) and one for PMCA4 (114 nt) (Strehler et al., 1989; Santiago-Garcia et al., 1996). In PMCA2 one additional insertion of 55 nt occurs, which is spliced at the 30 side of the 172 nt exon (Heim et al., 1992). Exclusion of all the exons generates the basic full length PMCA proteins designated as variant b, but all site C insertions generate a shift of the reading frame creating a premature stop codon. This truncates the protein, eliminating about half of the calmodulin-binding domain. Splicing events including the internal donor splice sites that generate insertions of 114 (PMCA1 and 4) and 87 nt (only in PMCA1) do not lead to the truncation of the pump, but modify the calmodulin-binding domain, and thus, the affinity of the pump for calmodulin. An additional 68 nt insertion has been found in the muscle isoform of PMCA3, corresponding to exon 22 of the rat

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. Figure 32‐2 Splicing variants of the four basic isoforms of the PMCA pump. Alternatively spliced exons are shown as gray, light-gray, and dark-gray boxes for splice site A, as light-gray and gray boxes for site C; flanking exons are shown as open boxes. The size of alternatively spliced exons is given in nucleotide numbers. A rat PMCA2 gene has been used as an example of the organization of alternatively spliced exons

PMCA3 gene. This exon contains an in frame stop codon, which leads to the generation of the shortest variant of the PMCA3 pump (Burk and Shull, 1992).

6.1 Functional Diversity of the PMCAs: Structure–Function Relationships The physiological rationale for the elevated number of PMCA pump variants is obscure. Very likely, there is no redundancy, each variant playing a specific role, or operating in specific sites of the cell. PMCA1 and PMCA4 appear to be less active than the neuronal PMCA2 and PMCA3 isoforms (Brini et al., 2003; Domi et al., 2007). The introduction of exons at site C, within the calmodulin-binding site, results in truncated

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variants (a or CII) that have lower affinity to calmodulin, at least in vitro (Hilfiker et al., 1994; Preiano et al., 1996). These truncated variants, however, generally have higher baseline ATPase activity in the absence of calmodulin (Preiano et al., 1996; Caride et al., 2001). The shorter variant of PMCA4 is more active in the native cellular environment than the full length variant (Brini et al., 2003) due to its faster kinetic of activation (Caride et al., 2007). As for the PMCA3 variants spliced at site C, both PMCA3b (CI) and PCMA3a (CII) surprisingly have similar activity when overexpressed in CHO cells (Brini et al., 2003). Somehow, even if truncated, the PMCA3a (CII) variant appears to retain the ability to react to calmodulin. The splicing events at site A, which lies within the catalytic domain in the intracellular loop between the 2nd and 3rd transmembrane domains, is particularly interesting in the PMCA2 isoform. As mentioned, it generates four variants. A study of the activity of the shortest (z) (AI) and the longest (w) (AIII) variants and that of the full length (b) (CI) and truncated (a) (CII) splicing at site C variants has revealed that the combination w/a of the PMCA2 has lower activity than all other PMCA2 splice variant combinations (w/b, z/a, and z/b) (Domi et al., 2007; Ficarella et al., 2007). This finding is of particular interest, since variant w is specifically sorted to the apical pole of polarized cells, where its reduced activity may have physiological meaning (Chicka and Strehler, 2003). The w/a, for instance, is the variant found in the stereocilia of the hair cells of the inner ear (Hill et al., 2006).

7

Regulation of the Activity of the PMCA Interaction with Modulators

The regulation of the PMCA activity by calmodulin was discovered in 1977 (Gopinath and Vincenzi, 1977; Jarrett and Penniston, 1977). As mentioned, calmodulin interacts with the calmodulin-binding domain (CBD) in the C-terminal tail of the PMCA protein (see > Figure 32‐3) and decreases its Kd for Ca2+ from 10–20 mM to Figure 32‐3). A growing body of evidence now indicates that this may indeed occur. For instance, the PMCAs, in particular PMCA4b, could be localized in the caveolae (Fujimoto, 1993; Hammes et al., 1998), which contain a number of effectors of cellular signaling pathways. All full length variants of PMCAs contain a C-terminal PDZ-binding domain, which mediates the interaction with members of the MembraneAssociated Guanylate Kinase family (MAGUK or SAP). These kinases contain a PDZ (PSD-95) domain, and have the role of assuring the interaction between ion transporters and other signaling proteins (Kim et al., 1998; DeMarco and Strehler, 2001). A member of the MAGUK family, the calcium/calmodulindependent serine protein kinase (CASK), was recently found to interact with PMCA4b (Schuh et al., 2003). This interaction is of particular interest, as in adult brain CASK is located at neuronal synapses, where PMCAs could be involved in the regulation of synaptic activity. Another PDZ domain-containing protein, Ania-3/Homer, which couples the NMDA-binding ionotropic glutamate receptors to metabotropic glutamatergic receptors (mGluR) and thus has a function in the regulation of synaptic activity (Sheng and Pak, 2000; Blackstone and Sheng, 2002), interacts with full length variants of all four PMCA isoforms (PMCA1b, 2b, 3b, 4b). The ubiquitous protein PISP also interacts with the b variants of all four isoforms. The interaction has a transient character and may play a role in sorting the PMCAs to or from the plasma membrane (Goellner et al., 2003). Proteins that contain PDZ domains interact with the full length variants of all PMCA isoforms, but others exhibit instead isoform selectivity. Thus, PMCA4b, but not PMCA2b, interacts through its C-terminal tail with the PDZ domain of inducible nitric oxide synthase 1 (NOS-1) (Schuh et al., 2001). The suggestion has been put forward that the activation of PMCA4b reduces the calcium concentration in the proximity to NOS-I, leading to its downregulation. A ternary complex composed of the PMCA (variants 1b or 4b), alpha-1 syntrophin and NOS-1 has been detected in cardiac myocytes (Williams et al., 2006). By contrast, PMCA2b, but not PMCA4b, interacts preferentially with the regulatory factor 2 of the Na+/H+ exchanger (NHERF2) leading to the functional assembly of PMCA2b in a multiprotein Ca2+ signaling complex, which is claimed to facilitate the cross-talk between local Ca2+ influx and efflux (DeMarco et al., 2002). The reasons for the isoform specificity in the interaction with NOS-1 and NHERF2 may be related to sequence differences in the C-terminal domains of the PMCAs. Studies searching for partners interacting with the main intracellular loop of the PMCA (between TM4 and TM5) have identified the catalytic subunit of calcineurin A (Buch et al., 2005) suggesting that PMCA

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could be a negative modulator of calcineurin-mediated signaling pathways. The proapoptotic tumor suppressor Ras-associated factor 1 (RASSF1) also interacts with the main intracellular region (652–748) of PMCA4b (Armesilla et al., 2004). Since this region has a high level of homology (about 90%) among the PMCA isoforms, this interaction may represent a general feature of the PMCAs. As mentioned, the N-terminal tail of PMCA has very low sequence homology among isoforms. A search for isoform-specific interactors with this region has identified protein 14–3–3, showing that isoform e specifically interacts with the PMCA1, 3, and 4 but not with the PMCA2, isoform resulting in the inhibition of pump activity (Rimessi et al., 2005; Linde, 2008).

8

PMCA Pumps and Pathology

8.1 Studies on PMCA Knock-Out Mice Knock-out (KO) mice have been produced for PMCA1, PMCA2, and PMCA4. The phenotypes have distinctive features. No KO mice have so far been produced for PMCA3. The broad tissue expression of PMCA3 mRNA between 12.5 and 15.5 days of gestation, which indicates a possible critical role for PMCA3 in normal embryonic development, may explain the failure. Homozygous PMCA1 KO mutants (Okunade et al., 2004) survived only until day 8.5 of gestation and then died, indicating that the presence of PMCA1 is necessary during embryogenesis. This would be consistent with its major housekeeping function. Somewhat surprisingly, however, the heterozygous mutants did not exhibit a pathological phenotype, even if the smooth muscle cells of the portal vein were prone to apoptosis. The PMCA4 KO mutants survived well. The homozygous mice appeared outwardly healthy and had no histopathological defects (Okunade et al., 2004). However, the null mutants exhibited male infertility, due to the inability of the sperm to achieve hyperactivated motility. The defect prevented it from traversing the female genital tract to fertilize the egg (Okunade et al., 2004; Schuh et al., 2004). PMCA4 accounts for more than 90% of the total PMCA protein in the testes: evidently, its loss cannot be compensated by other PMCA isoforms. The homozygous PMCA2 mutant mice survived, but grew more slowly than the heterozygous and wild-type littermates. However, the null mutants exhibited severe ataxia by about day 12 of age, with an unsteady gait and difficulty in maintaining balance (Kozel et al., 1998). Histological analysis of the inner ear revealed the absence of otoconia and a number of alterations in the organ of Corti. Unexpectedly the cerebellum maintained normal gross organization and cell distribution, even if minor alterations were present: e.g., decreased number of granular neurons, increased number of Purkinje cells, and decreased thickness of the molecular layer. Thus, the balance impairment evidently was due to the absence of otoconia that sense changes in equilibrium, and not to cerebellar neuronal impairment. Auditory brainstem responses showed that the KO mice were profoundly deaf, while heterozygous mice had significant hearing loss. Evidently, the PMCA2 pump is essential for the hearing process.

8.2 PMCA2 Mutants and Hereditary Hearing Loss A point mutation of PMCA2 was reported in a mouse termed deafwaddler (dfw), affected by hereditary hearing loss (Street et al., 1998). In the mutant, an A to G nucleotide transition in DNA caused a G283S substitution at a highly conserved amino-acid position in the first intracellular loop (between TM2 and TM3) of the pump (> Figure 32‐4). The Dfw mutant was deaf and displayed vestibular/motor imbalance. In homozygotes, the mutation caused a number of ultrastructural changes in the inner ear, including apoptosis of outer hair cells, disappearance of the stereocilia from the second and third rows of outer hair cells, and loss of hair cells and the spiral ganglion cells in older mice (Dodson and Charalabapoulou, 2001). Another PMCA2 mutation was described in a second allele, dfw2J. This was a 2-base-pair deletion, which caused a frame-shift and a truncated protein, and led to its disappearance from the stereocilia and the

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. Figure 32‐4 Deafness-associated mutations of PMCA2 pump. Deafwaddler mice bear a G283S mutation in the first intracellular loop of the pump. A G293S substitution has been identified in human family: the mutation in the PMCA2 gene was accompanied by a CDH23 point mutation, causing a digenic mechanism of hearing loss. The Wriggle Sagami mouse carries a K412E mutation in the N-terminal portion of TM4. A V586M substitution in the ATP binding site associated with a mutation in CDH23 gene has been found to increase hearing impairment in three of five siblings of a human family. In the left panel: sequence alignment showing the mutation sites in the PMCA2 gene shown in the right panel. Conserved amino acid substitutions are highlighted, and the number of the mutated residue is given. Note the conservation of the affected residues in mammalian species

baso-lateral membrane of cochlear hair cells (Street et al., 1998). A third mutation has also been described: the Wriggle Mouse Sagami carried an E412K substitution in the fourth transmembrane domain of the PMCA2 pump (Takahashi and Kitamura, 1999) (> Figure 32‐4). The homozygous wri mutants exhibit neurological pathology with motor imbalance and progressive hearing loss. Recently, two PMCA2 mutations leading to hearing loss in humans have also been described (Schultz et al., 2005; Ficarella et al., 2007). In both cases the PMCA2 mutation was associated with mutations in the cadherin-23 gene (CDH23). Schultz et al. (Schultz et al., 2005) have described five siblings carrying a homozygous point mutation of CDH23 and exhibiting recessive sensorineural hearing loss. Three of five siblings had an additional heterozygous substitution in PMCA2 (V586M), producing a hypofunctional pump variant associated with increased hearing loss (Schultz et al., 2005) (> Figure 32‐4). Ficarella et al. have recently identified another PMCA2 mutation (G293S) that, in association with a point mutation in the CDH23 gene (T1999S), caused digenic hearing loss (Ficarella et al., 2007) (> Figure 32‐4). Analysis of the activity of the recombinant (G293S) PMCA2 mutant overexpressed in CHO cells showed that the mutant had a dramatic defect in the calcium handling by the pump, i.e., the pump had reduced ability to clear Ca2+ from the stereocilia (Ficarella et al., 2007).

8.3 PMCA Pumps in Other Pathological Conditions Apart from hereditary hearing loss, no human genetic disease has so far been associated with a genetic alteration of the PMCA genes. However, the function of the PMCA pump has been found altered in several pathological conditions of the nervous system, including ageing and ischemia/anoxia.

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Calcium homeostasis is often altered in neurodegenerative diseases. The amyloid-beta peptide (A-beta), which accumulates abnormally in the neurons of Alzheimer’s disease patients, causes disturbances in calcium homeostasis in hippocampal neurons (Mark et al., 1995). Downregulation of the PMCA2 isoform at both the transcriptional and protein levels was observed during experimental autoimmune encephalomyelitis (EAE) (Nicot et al., 2003). The PMCA2 pump deficiency was apparently causative, highlighting the importance of PMCA2 in the neuronal dysfunctions linked to inflammation and neurodegeneration (Kurnellas et al., 2005; Nicot et al., 2005). Abnormalities in cardiac function and in the regulation of blood pressure may be linked to alterations of PMCA pumps. Sarcolemmal membranes from rats with cardiac hypertrophy exhibited increased PMCA pump activity (Nakanishi et al., 1989). The alteration may represent an adaptive mechanism to facilitate removal of Ca2+ from the myocardial cells during the development of cardiac hypertrophy (Nakanishi et al., 1989). In platelets of hypertensive individuals, instead, PMCA activity was inhibited, resulting in increased intracellular Ca2+, in hyperactive platelets, and in increased risk of heart attack and stroke (Blankenship et al., 2000). Nitric oxide produced by nNOS is now gaining attention as a signaling molecule active in the regulation of excitation–contraction (EC) coupling, cell growth, and hypertrophy of myocardium (Cartwright et al., 2007; Takimoto and Kass, 2007). The functional interaction between nNOS and the PMCA pump is of interest, since PMCA isoforms 1 and 4 have been found to interact with nNOS in vitro and in vivo (Schuh et al., 2001; Williams et al., 2006). Overexpression of PMCA4 in HEK293 cells dramatically downregulated nNOS activity, whereas the expression of a variant of PMCA4 with a deleted PDZ-binding domain failed to do so (Schuh et al., 2001). In another model, mice overexpressing PMCA4 in arterial smooth muscles under the control of the SM22a promoter displayed higher blood pressure than control animals (Gros et al., 2003; Schuh et al., 2003). Downregulation of PMCA activity by reactive oxygen species (ROS), mediated by increased tyrosine phosphorylation, was found in platelets from patients with non-insulin-dependent diabetes mellitus (Rosado et al., 2004; Jardin et al., 2006). Finally, a growing body of evidence indicates the involvement of PMCA dysregulation in the pathogenesis of cancer (Monteith et al., 2007). Thus, specific upregulation of PMCA4, but not of PMCA1, has been found during the differentiation of the HT-29 colon cancer cell line (Aung et al., 2007). The levels of transcript coding for the PMCA2 pump increased strongly in some breast cancer cell lines (Lee et al., 2005a). Moreover, antisense-mediated downregulation of PMCA pumps dramatically reduced the proliferation of MCF-7 breast cancer cells (Lee et al., 2005).

9

Concluding Remarks

Forty years after its discovery the plasma membrane calcium pump has become an important topic in the booming area of Ca2+ signaling. Several aspects of this pump distinguish it clearly from the other two calcium pumps, but also make its study particularly difficult. Progress in recent years has shown that the pump has functions that are more diversified than those of the SERCA pump and the SPCA pump. This is related to the unusually large number of basic isoforms and alternative spliced variants of the PMCA pump: the area is now emerging of specific functions of the PMCA pump in different cell types and even in different domains of the cell. The PMCA pump is not only critical in the maintenance of the appropriately low cytosolic Ca2+ concentration: it also shapes the calcium signals spatially and temporally. The pump is essential to the normal functioning of cell life, and disease conditions are now beginning to appear, which can be traced back to the malfunction of the PMCA pump. This is particularly true of nervous cells, which demand an absolutely precise control of calcium homeostasis.

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Calcium and Apoptosis

J. Guo . Y. Lao . D. C. Chang

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 598

2 2.1 2.2 2.2.1 2.2.2 2.2.3

Apoptosis and the Apoptotic Signaling Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 599 Apoptosis and Other Forms of Cell Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 599 Apoptotic Signaling Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 600 Mitochondria Apoptotic Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 601 Death Receptor Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 601 Endoplasmic Reticulum (ER) Apoptotic Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 602

3 3.1 3.2 3.3 3.4

Role of Ca2+ Signaling in Programmed Cell Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 602 Whether Ca2+ Signal is Involved in Programmed Cell Death? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 602 Ca2+ Signaling is Involved in Regulating the Apoptotic Pathway Upstream of Mitochondria Dysfunction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 604 Downstream Targets of the Cytosolic Ca2+ Signal . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 604 The Crosstalk Between the ER and Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 608

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Bcl-2 Family Proteins and Ca2+ Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 609 Bcl-2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 609 Bax/Bak . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 611 Other Bcl-2 Family Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 611

5 5.1 5.2 5.3

Ca2+ Signaling and Neuronal Cell Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 611 Ca2+ and ROS-Induced Neuronal Cell Death . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 611 ROS-Regulated Plasma Membrane Ca2+ Signaling Pathways in Neuronal Cell Death . . . . . . . . . . 613 ROS-Regulated ER Ca2+ Signaling Pathways in Neuronal Cell Death . . . . . . . . . . . . . . . . . . . . . . . . . . . 614

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Conclusion and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 614

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Calcium and apoptosis

Abstract: Calcium ion is an important secondary messenger in controlling cell death. The role of calcium in apoptotic signaling has been extensively investigated in recent years. Our overall understanding on this topic, however, is still far from complete. In fact, results of some of the earlier studies were somewhat controversial. Only in recent years, the role of Ca2+ signaling in apoptosis has become clear. In this review, we summarize some of the major aspects of Ca2+ signaling in apoptosis based on our own studies as well as those reported in the recent literature. This review includes four major parts: In Part 1, we give an overview of cell death and a summary of the essential apoptotic signaling pathways. In Part 2, we discuss the role of Ca2+ signaling in programmed cell death. Particularly, we presented the evidence that Ca2+ signal plays a role in the upstream of the mitochondria-dependent apoptotic pathway. In Part 3, we summarize the recent evidence suggesting possible interactions between the Ca2+ signal and Bcl-2 family proteins, which are wellknown regulators of the apoptotic process. Finally, in Part 4, we summarize recent findings on some of the Ca2+-regulating proteins that provide possible molecular pathways connecting Ca2+ signaling with ROSinduced neuronal cell death. We hope that, a better understanding of the Ca2+ signaling pathways may lead to the development of novel therapeutic approaches for major neurodegenerative diseases. List of Abbreviations: AIF, apoptosis-inucing factor; Apaf-1, apoptotic protease activating factor-1; Bax, Bcl-2-associated X protein; Bcl-2, B cell lymphoma-2; tBid, truncated Bid; Bim, Bcl-2 interacting mediator; Cyt c, cytochrome c; DAP kinases, death-associated protein kinases; ER, endoplasmic reticulum; H2O2, hydrogen peroxide; IAP, inhibitor of apoptosis protein; IP3R, inositol 1,4,5-triphosphate receptor; PMCA, plasma membrane Ca2+ ATPases; PTP, permeability transition pore; ROS, reactive oxygen species; SERCA, sarco-endoplasmic reticulum Ca2+ ATPase; Smac, second mitochondria-derived activator of caspases; SR, sarcoplasmic reticulum; TNF-a, tumor necrosis factor-a; TRPM, transient receptor potential protein, melastatin subfamily; UPR, unfolded protein response; VDCC, voltage-dependent Ca2+ channel

1

Introduction

Apoptosis, also called programmed cell death, is a well-controlled cell suicidal process that allows organism to destroy damaged or unwanted cells in an orderly way (Kerr et al., 1972; Raff et al., 1993; Williams and Smith, 1993). Apoptosis is a very important cellular process that is essential for maintaining our normal physiological function. It has gained wide-spread attention in recent years because of its newly discovered roles in a variety of physiological and pathological processes. For example, when DNA is damaged in a cell and cannot be repaired, the cell will enter apoptosis to avoid the formation of abnormalities in the tissue. Thus, failure of programmed cell death can cause cancer. Besides tumorigenesis, malfunction of apoptosis is also found to be associated with many neurodegenerative diseases, including Alzheimer’s disease, Parkinson’s disease and stroke. Because of the physiological and pathological importance of apoptosis, its study has attracted tremendous scientific interests in the last ten years. Results from recent investigations have revealed that apoptosis is controlled by a complex network of signal transduction pathways. At this time, it is still not totally clear how apoptosis is precisely regulated by the external and internal signals, although a number of key regulators have been identified. These regulators include protein factors released from the inter-membrane space of mitochondria, the Bcl-2 family proteins, and different members of caspases. Since it is well known that overloading of Ca2+ is frequently associated with neuronal cell death, it was suspected that Ca2+ signal may play an important role in apoptosis. As one of the major second messengers, Ca2+ signal is known to be involved in regulating many important physiological functions. Thus, many studies were conducted in the past two decades to examine the functional role of Ca2+ signaling in apoptosis. Results of these studies were somewhat controversial. Some early studies reported that Ca2+ elevation was involved only in the later stage of cell death during DNA degradation, but played no significant role in initiating the apoptotic process. This view, however, was not shared by later studies. Indeed, since the beginning of this century, a large number of studies using different approaches have supported a general conclusion that Ca2+ signal is involved in the upstream apoptotic pathway in a variety of apoptotic models.

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One of the difficulties in studying the apoptotic signaling pathways is that apoptosis is not the only type of cell death. At present, three major types of cell death are known, including apoptosis, necrosis and autophagic cell death (Lockshin and Zakeri, 2004). The boundaries of these different types of cell death are sometimes overlapping. For example, when a neuronal cell is exposed to a certain dosage of ROS, it may start to undergo apoptosis first and then switch to necrosis later. Furthermore, although the term ‘‘programmed cell death’’ is generally referring to apoptosis, in some literatures it is also used to include both apoptosis and autophagic cell death.

2

Apoptosis and the Apoptotic Signaling Pathways

2.1 Apoptosis and Other Forms of Cell Death The term ‘‘apoptosis’’ was first introduced in 1972 by Kerr et al. (1972). Since then, this process has been studied extensively and was found to have an enormous importance in the development of biological organisms as well as the maintenance of their normal life. For example, apoptosis is utilized to eliminate the unwanted cells so that the proper structure of functional organs can be formed in the embryo. In fact, formation of digits in a hand and the proper targeting of the neural connection are known to be dependent on apoptosis. Thus, defects in apoptosis will result in major developmental abnormalities (Jacobson et al., 1997). Apoptosis is also used in the thymus to eliminate self-reactive T cells to avoid autoimmunity (Thompson, 1995). Furthermore, apoptosis is a major mechanism for preventing cancer (Fisher, 1994; Reed, 2003). When a cell has excessive DNA damage, it will trigger the apoptotic process to kill the damaged cell. If the apoptotic mechanism fails, it will result in tumorigenesis and/or the development of chemotherapy resistance. At present, every type of human cancer is known to be associated with defects in the apoptotic mechanism (Johnstone et al., 2002; Reed, 2003; Fesik, 2005). Thus, defective apoptosis can cause multiple pathological states, including developmental abnormalities, cancer, autoimmune diseases, and neurodegenerative diseases. Because of its importance, in recent years its study has attracted a very large group of scientists from many fields, including cell and developmental biology, physiology, immunology, cancer research, and neuroscience. Unlike the accidental death process such as necrosis, apoptosis is a highly controlled physiological process of removing cells within an organism without destroying the surrounding tissue or to cause an inflammation (Kerr et al., 1972). Cells undergoing apoptosis are characterized by several clear morphological changes, including shrinkage and blebbing of cells, condensation of their chromatin followed by internucleosomal cleavage of DNA into regular fragments, and turnover of phosphatidylserine to the outer leaflet of the plasma membrane. The cellular contents, after digested by a group of specific proteases called caspases, are packed into membrane-surrounding vesicles. These vesicles, called ‘‘apoptotic bodies,’’ are then taken up by neighboring cells through phagocytosis. In contrast to apoptosis, necrosis is a form of uncontrolled cell death, in which the cell swells and ruptures in response to profound damage. Typically, cells entering necrosis undergo lysis and release many intracellular constituents. As a result, this process usually provokes an inflammatory response (Nelson and White, 2004). There is evidence that some defined pathways may be involved in necrosis (Yuan et al., 2003). The distinction between apoptosis and necrosis may be related to the timing of the response and the severity of insult (Lockshin and Zakeri, 2004). Some common signaling pathways, such as death receptors, kinase cascades, and mitochondria, appear to participate in both processes. Hence, by modulating these pathways, it is possible to switch between apoptosis and necrosis. Another form of cell death, autophagy (from the Greek, ‘‘self eating’’), enables eukaryotic cells to capture cytoplasmic components for degradation within lysosomes. It proceeds through a sequence of morphological changes in a highly regulated process. Briefly, the autophagic pathway begins with the sequestration of cytoplasmic material in double-membrane vesicles known as autophagosomes (Klionsky and Emr, 2000). The sequestration process is under the control of GTPases (Ogier-Denis et al., 1996) and involves novel ubiquitin-like conjugation systems (Ohsumi, 2001). The autophagosome then fuses with the

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lysosome to form an autolysosome. The inner vesicle is degraded by resident hydrolases together with its cargoes (Klionsky and Emr, 2000). Recent studies in mammalian cells reveal that autophagy is controlled by complex signaling pathways. In these pathways, phosphatidylinositol 3-kinase (PI3K) is required to generate autophagosomes and a negative regulator of autophagy is the target of rapamycin protein (TOR) kinase (Meijer and Codogno, 2006).

2.2 Apoptotic Signaling Pathways Apoptosis is controlled by a very complicated network of signal transduction pathways. At present, the detailed mechanisms for regulating apoptosis by various external and internal clues are still not totally understood. In general, three major signaling pathways have been identified in apoptosis (Strasser et al., 2000) (> Figure 33-1). They are: (1) the mitochondrial pathway (Desagher and Martinou, 2000; Wang, 2001), (2) the death receptor pathway (Ashkenazi and Dixit, 1998; Sun et al., 1999), and (3) the endoplasmic reticulum (ER) pathway (Nakamura et al., 2000). All of these pathways finally converge to the activation of the executioner caspase-caspase 3, which destroys various subcellular components by proteolysis (Cohen, 1997).

. Figure 33-1 A schematic diagram showing the three major signaling pathways in apoptosis, i.e., the death receptor pathway, the mitochondria-dependent pathway and the ER pathway (For details, see text)

Calcium and apoptosis

33

2.2.1 Mitochondria Apoptotic Pathway Mitochondrial pathway is the most commonly used mechanism in animal cells to execute apoptosis (Green and Reed, 1998; Ravagnan et al., 2002). In this pathway, mitochondria play a central role in regulating the progression of apoptosis (> Figure 33-1). Briefly, the extracellular or intracellular apoptotic signals trigger the activation of the pro-apoptotic Bcl-2 family proteins, such as Bax, in the cytosol. These apoptotic regulators translocate to mitochondria and induce the release of apoptotic factors [including cytochrome c, Smac (second mitochondria-derived activator of caspases), AIF (apoptosis inducing factor) and endonuclease G, etc.] from the inter-membrane space of mitochondria (Kluck et al., 1997; Reed, 1997; Wang, 2001). One of the key apoptotic factors released is cytochrome c (cyt c), which binds to Apaf-1 (apoptotic protease activating factor-1) and activates Apaf-1 in the presence of dATP (Zou et al., 1997). The activated Apaf-1 then recruits procaspase-9 and triggers its activation by auto-cleavage. The activated caspase-9 in turn cleaves procaspase-3 and generates an active tetrameric form (Li et al., 1997). The catalytic sites of the processed caspases, however, are immediately bounded by the IAPs (inhibitor of apoptosis proteins) (Riedl et al., 2001). During apoptosis, another apoptotic factor, Smac/DABLO, is co-released from mitochondria (Du et al., 2000; Verhagen et al., 2000). Smac can bind with IAPs and remove them from the processed caspases, and thus allow the caspases to be catalytically active (Verhagen and Vaux, 2002). Although the downstream events in the mitochondrial pathway are relatively well known, the upstream events are far less clear. It is only known that Bcl-2 family proteins play a key regulatory role in permeabilizing the mitochondrial outer membrane. The Bcl-2 family proteins contain four conserved Bcl-2 homology (BH) domains designated as BH1, BH2, BH3, and BH4. On the basis of the structural and functional characteristics, members of Bcl-2 family proteins can be divided into two groups as anti-apoptotic proteins (such as Bcl-2 and Mcl-1) and pro-apoptotic proteins (such as Bax and Bim). The pro-apoptotic proteins can be subdivided into two categories: those containing BH1, BH2, and BH3 domains (such as Bax and Bak), called ‘‘multidomain pro-apoptotic proteins’’ (Gross et al., 1999) and those containing only the conserved BH3 domain (such as Bim), called ‘‘BH3-only proteins’’ (Strasser, 2005). The detailed mechanisms on how these proteins regulate apoptosis are still under active investigation. It is generally thought that, under an apoptotic stress signal, pro-apoptotic proteins such as Bax and Bak can form pores in the mitochondrial outer membrane, resulting in releasing the apoptotic factors in the inter-membrane space (Korsmeyer et al., 2000; Scorrano and Korsmeyer, 2003). Activation of these proapoptotic proteins is likely to be caused by transient interaction with the BH3-only proteins (Korsmeyer et al., 2000). On the other hand, the anti-apoptotic proteins such as Bcl-2, Mcl-1 and Bcl-xL can neutralize the BH3-only and pro-apoptotic proteins, and prevent them to form pores in the mitochondria.

2.2.2 Death Receptor Pathway The death receptor pathway is initiated by extracellular death-inducing ligands of the TNF (tumor necrosis factor) superfamily such as TNF-a, FasL/CD95L, TWEAK (TNF-like weak inducer of apoptosis) and TRAIL (tumor necrosis factor-related apoptosis-inducing ligand) (Krammer, 2000; Screaton and Xu, 2000; Locksley et al., 2001). These ligands bind to cell surface death receptors (TNFR, Fas/CD95, DR3, DR4/DR5 respectively) leading to their trimerization. Then, the trimerized death receptors recruit cytoplasmic adaptor proteins such as TRADD (TNF receptror-associated death domain) and/or FADD (Fas-associated death domain) via a death domain (DD). These adaptors form a death-inducing signaling complex (DISC) with the initiator caspase-8 via another interaction motif, the death effector domain (DED). Procaspase-8 is autoactivated at DISC and becomes the active enzyme caspase-8, which is released into the cytosol. Once activated, caspase-8 cleaves and activates downstream effector caspases such as caspase-3, caspase-6 and caspase-7 to amplify the apoptotic death signal. Caspase-8 can also activate Bid, a BH3-only pro-apoptotic Bcl-2 family protein, by cleavage. The truncated Bid (tBid) then translocates to mitochondria and activates the mitochondria-dependent apoptotic pathway (Luo et al., 1998).

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2.2.3 Endoplasmic Reticulum (ER) Apoptotic Pathway In the ER pathway, caspase-3 is activated through an ER-located caspase, caspase-12 (Nakagawa et al., 2000). Caspase-12 is predominately situated at the cytosolic side of the ER membrane, and is specifically activated by disturbance to ER homeostasis, such as ER Ca2+ depletion. The activation of caspase-12 has been shown to depend on a Ca2+-dependent protease calpain (Nakagawa and Yuan, 2000). In addition, the ER can regulate the apoptotic signaling through IP3 receptor (IP3R) or ryanodine receptor (RyR) mediated by Ca2+ release. Another important ER stress signal is the unfolded protein response (UPR). The efficient functioning of the ER is essential for cell survival. Conditions that interfere with ER function lead to the accumulation and aggregation of unfolded proteins. There are specific receptors in the ER detecting the onset of ER stress and initiate the unfolded protein response to restore normal ER function. If the stress is prolonged, or the adaptive response fails, the cell will enter apoptosis (Szegezdi et al., 2006). ER stress-induced apoptosis has been implicated in the pathophysiology of several neurodegenerative and cardiovascular diseases (Marciniak and Ron, 2006; Yoshida, 2007).

3

Role of Ca2+ Signaling in Programmed Cell Death

3.1 Whether Ca2+ Signal is Involved in Programmed Cell Death? Ca2+ ions are recognized as a major intracellular messenger that regulates many cell and tissue physiological processes. Intracellular Ca2+ changes are also implicated in a number of pathological states such as cardiac ischemia (Treiman, 2002; Hernandez et al., 2003), muscular dystrophy (Gailly, 2002), neuronal damage during cerebral ischemia (Paschen, 2003; Verkhratsky and Toescu, 2003), and hypoglycemia (Dworakowska and Dolowy, 2000). The involvement of Ca2+ in cell death has been suggested almost as early as the discovery of apoptosis. In 1974, it was shown that excess Ca2+ entry into cardiac myofibers causes cell demise, playing a pathological role in cardiac myopathies (Fleckenstein et al., 1974). The functional connection between [Ca2+]c increase and cell death was further studied by examining the effects of either artificially increasing [Ca2+]c with a Ca2+ ionophore A23187 or suppressing [Ca2+]c elevation with a Ca2+ chelator on cell death. It was found that A23187 could mimic the apoptotic effect of glucocorticoid; while chelating Ca2+ in the medium with EGTA could block glucocorticoid-induced cell death (Kaiser and Edelman, 1977). McConkey et al. have carried out a series of works to demonstrate that endonuclease activation and cell death were dependent on a sustained increase in cytosolic Ca2+ concentration in a variety of apoptotic inducers and cell models (McConkey et al., 1988, 1989a–c, 1991, 2000; McConkey, 1996). Similar results were obtained later from studies in many other apoptotic systems (Aw et al., 1990; Perotti et al., 1990; Chow et al., 1992; Zhivotovsky et al., 1993, 1994; Szondy, 1994; Marin et al., 1996; Kobayashi et al., 1997; Kimura et al., 1998). In addition to Ca2+ elevation in cytosol, it was suggested that Ca2+ depletion from the ER could also play an important role in the initiation of apoptosis (Nakamura et al., 2000; Pan et al., 2000). For example, cyt c release was observed in cells treated with thapsigargin, an irreversible inhibitor of the Ca2+-ATPase in ER membrane, suggesting that ER Ca2+ depletion could be a triggering mechanism for the early apoptotic event of cyt c release. Although these earlier studies strongly suggest that Ca2+ may play a significant role in the signaling pathway of apoptosis, several important issues, however, are yet to be resolved. First, results of the ionophore experiments only indicate that a high level of [Ca2+]c is a death inducer; they do not necessarily imply that elevation of [Ca2+]c is required in the normal apoptotic process induced by other treatments. Second, it is still not clear whether the observed [Ca2+]c elevation is the result or the cause of certain apoptotic events. In order to demonstrate more convincingly the involvement of Ca2+ signal in regulating apoptosis, one must show that apoptosis can be prevented by blocking [Ca2+]c elevation. A number of research groups had conducted such Ca2+ blocker experiments using a membranepermeable form of Ca2+ chelator (BAPTA/AM) to suppress the Ca2+ signal within apoptotic cells. The results, however, were very controversial. While some studies indicated that BAPTA/AM could suppress

Calcium and apoptosis

33

apoptosis to a certain extend (Kruman et al., 1998; Shan et al., 1998; Shi et al., 1998; Ko et al., 2000), others reported that BAPTA/AM could not prevent apoptosis at all (Hampton et al., 1996; Voehringer et al., 1997; Wei et al., 1998; Zhou et al., 1998; Wertz and Dixit, 2000; McFarlane et al., 2000; Pan et al., 2000). In some cases, it was found that treatment of BAPTA/AM itself could induce DNA fragmentation and cell death (Grant et al., 1995). Taken together, the above results raised a serious question on whether Ca2+ signaling is truly involved in regulating the progression of apoptosis. In order to clarify the question of whether cytosolic Ca2+ signal is involved in regulating the progression of apoptosis, we have conducted a series of experiments on the effects of three different types of Ca2+ signal blocker on UV- or H2O2-induced apoptosis (Pu and Chang, 2001). We found that the membrane-permeable form BAPTA/AM could only inhibit H2O2-induced apoptosis in neuroblastoma cells (unpublished data), but not in UV-induced apoptosis in Hela cells (> Figure 33-2a). However, the membrane-impermeable form BAPTA could inhibit UV-induced apoptosis in Hela cells (> Figure 33-2b).

. Figure 33-2 The effect of BAPTA/AM, BAPTA and heparin on UV-induced apoptosis in HeLa cells. (a) The effect of membrane-impermeant BAPTA/AM on UV-induced apoptosis. (b) The effect of membrane-permeant BAPTA/AM on UV-induced apoptosis. (c) The effect of IP3R blocker Heparin on UV-induced apoptosis. (Reproduced from Pu and Chang, 2001)

Results of our studies demonstrated clearly that by blocking the increase of [Ca2+]c, a significant percentage of cells were prevented to enter apoptosis in both systems. Blocking the release of Ca2+ from the ER by applying an IP3R antagonist (heparin) had a similar effect (> Figure 33-2c). These findings strongly suggest that cytosolic [Ca2+] increase is involved in the signaling pathway that drives the progression of apoptosis. Then, how can one explain the controversial results of BAPTA/AM? We think that the cause of diverse effects of BAPTA/AM mentioned above could be due to the dual effect of BAPTA/AM on

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sub-cellular Ca2+ distribution. Since BAPTA/AM is membrane permeable, it can center the cytoplasm as well as subcellular organelles such as the ER in some cell types and accumulate there. Thus, BAPTA/AM would not only block the elevation of [Ca2+]c, it could also chelate the free Ca2+ in the ER lumen. Since depletion of Ca2+ in ER could stimulate apoptosis (Luo et al., 1998; Sidoti-de Fraisse et al., 1998; Verhagen et al., 2000; Du et al., 2000), BAPTA/AM could have an unwanted side effect that would enhance programmed cell death in some apoptotic models.

3.2 Ca2+ Signaling is Involved in Regulating the Apoptotic Pathway Upstream of Mitochondria Dysfunction The process of apoptosis involves two major stages: the early ‘‘commitment phase’’ and the later ‘‘execution phase.’’ It had been reported that a sustained [Ca2+]c elevation was observed during the execution phase of apoptosis (Perotti et al., 1990; Zhivotovsky et al., 1994; McConkey et al., 2000). This sustained [Ca2+]c elevation was believed to be involved in the triggering mechanism of DNA fragmentation (McConkey et al., 1989b; McConkey and Orrenius, 1997). A more important question is whether Ca2+ is involved in regulating apoptosis in the commitment phase. During the 1990s, there was a very diverse view on this question. Some studies suggested that Ca2+ may interact with early apoptotic events. For example, an elevated [Ca2+]c could cause isolated mitochondria to swell and release their cyt c (Andreyev and Fiskum, 1999; Gogvadze et al., 2001; Schild et al., 2001). Moreover, T cells deficient in IP3 receptor were resistant to apoptosis in response to dexamethasone, ionizing radiation, or Fas stimulation (Jayaraman and Marks, 1997, 2000). Other groups, however, thought that Ca2+ signal may not be involved in the early phase of apoptosis. For example, in a ratio imaging measurement using fura-2 on apoptotic HL-60 cells, Lennon et al. observed an increase of [Ca2+]c at the later stage of apoptosis. According to their interpretation, these increases were apparently a result of, rather than a cause of, apoptosis (Lennon et al., 1992). Using a Ca2+ fluorescent probe Fluo-3, Scoltock et al. detected a sustained [Ca2+]c elevation between 1 and 2 hr after antiFas treatment in Jurkat cells. They attributed this [Ca2+]c increase to be only a requirement for DNA degradation, and not for an early apoptotic event (Scoltock et al., 2000). In order to resolve this question in a convincing way, one needs to conduct at least two direct experiments. First, one must directly measure the dynamic variation of cytosolic Ca2+ to show [Ca2+]c changes during an early stage of apoptosis; second, one needs to apply specific Ca2+ signaling blockers to show that apoptosis can be blocked at a stage upstream of cyt c release. Using a sensitive fluorescence probe of Ca2+ (calcium green-1 conjugated with 10 kD dextran), our laboratory had detected cytosolic [Ca2+] changes during the process of apoptosis in two apoptotic models, UV- and TNFa-induced apoptosis in HeLa cells (Pu et al., 2002). In our model, we observed a series of repetitive Ca2+ spikes in the early stage of apoptosis in many UV- or TNFa- treated Hela cells (> Figure 33-3). We found that these transient [Ca2+]c spikes occurred in a time window before cyt c release (Pu et al., 2002). Furthermore, when we blocked the [Ca2+]c increase by injecting BAPTA or heparin into the cells, cyt c release was effectively prevented or delayed (> Figure 33-4). These results strongly suggest that an early Ca2+ signal plays a role in regulating the progression of apoptosis upstream of cyt c release from mitochondria (Pu et al., 2002).

3.3 Downstream Targets of the Cytosolic Ca2+ Signal Elevation of [Ca2+]c may result in activation of a wide variety of signaling transduction pathways. However, it still remains elusive onhow Ca2+ elevation may be involved in apoptosis induction. Two major signaling pathways have been proposed (Rizzuto et al., 2003). In one scenario, cytosolic Ca2+ elevation can lead to the activation of many Ca2+-dependent enzymes, including calpain and calcineurin, which in turn drive the apoptosis progression by activating downstream proteins. Alternatively, mitochondrial Ca2+ overloading in response to the cytosolic Ca2+ increase could trigger a change in the mitochondrial permeability transition pores (PTP) and cause mitochondria dysfunction (see > Section 3.4). > Figure 33-5 summarizes the possible mechanisms of Ca2+ signaling in regulating the progression of apoptosis.

Calcium and apoptosis

33

. Figure 33-3 Dynamic changes of [Ca2+]c in HeLa cells during apoptosis. (a) Time-dependent changes of [Ca2+]c in HeLa cells undergoing UV-induced apoptosis as indicated by a Ca2+ imaging measurement using Calcium green-1 (CaGn-1). (b) Time-dependent changes of [Ca2+]c in HeLa cells undergoing TNF-a-induced apoptosis. (c) Correlation between the observed [Ca2+]c elevation and the change of cell size in a HeLa cells undergoing UV-induced apoptosis during the commitment phase and the execution phase

Calpain. Many studies suggested that calpain may work downstream of the cytosolic Ca2+ increase to activate different apoptotic players: (1) Calpain may activate Bax by cleavage (Wood et al., 1998). N-terminal cleavage of Bax by calpain generates a potent proapoptotic 18 kD fragment that promotes cyt c release (Gao and Dou, 2000; Wood and Newcomb, 2000; Choi et al., 2001). (2) Calpain can activate Bid

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. Figure 33-4 BAPTA or heparin can block UV-induced cyt c release in HeLa cells. (a) Immunostaining shows the filamentous distribution pattern of endogenous cyt c in control cells without apoptotic treatment. (b) Immunostaining shows the diffused distribution pattern of cyt c in a HeLa cell undergoing UV-induced apoptosis. Cyt c had already been released from mitochondria to cytosol in this cell. (c) Percentage of cyt c-released cells during UV-induced apoptosis in the presence (or absence) of BAPTA or heparin. The value presented in each bar is the average of four independent experiments, in each of which more than 1,000 cells were analyzed. (Reproduced from Pu et al., 2002)

by cleavage. It has been shown that in human melanoma cells the apoptotic drug cisplatin induced cleavage of Bid, yielding a 14 kD fragment, which could trigger cyt c release in isolated mitochondria (Mandic et al., 2002). Chen et al. also demonstrated in vitro that the recombinant Bid was cleaved by calpain into a fragment that was able to trigger cyt c release (Chen et al., 2001). (3) The activation of the ER-located caspase, caspase-12, may be dependent on calpain activity. It has been shown that calpain inhibitors could prevent caspase-12 activation in ER stress-induced apoptosis (Nakagawa and Yuan, 2000). (4) Calpain could abolish the inhibitory effect of XIAP (X-linked inhibitor of apoptosis protein) on caspases. Kobayashi et al. showed recently that calpain cleaved XIAP in vitro, producing fragments that are unable to bind the catalytic site of caspase-3 (Kobayashi et al., 2002). (5) Calpain could facilitate the progression of apoptosis through activating the calcineurin-dependent pathway. It has been shown recently that calpain mediated calcium-triggered apoptosis through cleaving the endogenous inhibitor of calcineurin, cain/cabin1. The cleavage of cain/cabin1 releases calcineurin from inhibition and activates the downstream calcineurindependent apoptotic pathway (Kim et al., 2002). In our study, we also observed that applying calpain inhibitors could partially suppress UV-induced apoptosis in Hela cells (unpublished results).

Calcium and apoptosis

33

. Figure 33-5 Possible mechanisms of Ca2+ signaling in regulating the progression of apoptosis. Upon apoptotic stimulation, an increase of free Ca2+ in the cytosol is generated via Ca2+ release from the ER or Ca2+ influxes from the extracellular medium. These excessive Ca2+ ions can bind to Ca2+-dependent enzymes, such as calpain and calcineurin, and activate their downstream targets including the Bcl-2 family proteins. Alternatively, the released Ca2+ may be directly taken up by mitochondria and such excessive Ca2+ uploading could trigger the mitochondrial dysfunction. The dashed arrows denote suggestive pathways that are not yet confirmed. (Reproduced from Guo et al., 2005)

Calcineurin. The involvement of calcineurin in apoptosis was suggested by the studies showing that cyclosporine A, an inhibitor of calcineurin, could block apoptosis in some apoptotic systems (Makrigiannis et al., 1994; Waring and Beaver, 1996). It was demonstrated that active calcineurin could dephosphorylate Bad (Wang et al., 1999). The dephosphorylated Bad is then released from its quenching protein 14-3-3, and translocates to mitochondria to trigger cyt c release (Wang et al., 1999). We also observed that calcineurin inhibitor could suppress UV-induced apoptosis in Hela cells (unpublished results). DAP kinases. DAP kinases are a family of death-associated protein kinases (Cohen et al., 1997; Cohen and Kimchi, 2001). Their regulatory mechanism is very similar to that of the Ca2+/calmodulin-dependent protein kinases II (CaMKII), with an auto-inhibitory CaM binding domain. In addition, DAP kinases also have an auto-inhibitory death domain at its C-terminal (Cohen and Kimchi, 2001). Like CaMKII, the activation of DAP kinases depends on transient Ca2+ elevation in the cytosol (Cohen et al., 1997; Cohen and Kimchi, 2001). The involvement of DAP kinases in TNF-a/Fas-induced apoptosis has been reported before (Cohen et al., 1999). However, the mechanism of how DAP kinases regulate the apoptotic process is still unknown.

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Nitric oxide synthase. The role of the nitric oxide synthase (NOS) in apoptosis is complex. NOS isozymes are constitutively expressed in endothelial cells (eNOS) and neurons (nNOS). Excessive production of nitric oxide (NO) following glutamate-stimulated Ca2+ entry through NMDA (N-methyl-Daspartate) receptors was implicated in excitotoxicity in cortical neuronal cultures (Dawson et al., 1991) and in brain ischaemia (Huang et al., 1994). The production of NO and superoxide anion result in the formation of peroxynitrite, which can diffuse from mitochondria to damage various cellular constituents. On the basis of the studies using selective nNOS antagonists (O’Neill et al., 2000) and nNOS knockout mice (Huang et al., 1994), it was shown that neuronal production of NO contributes to ischemic cell death.

3.4 The Crosstalk Between the ER and Mitochondria Recently, mitochondrial Ca2+ overloading has been shown to play a crucial role in apoptosis induction (Szalai et al., 1999; Pinton et al., 2001). What is the Ca2+ source from which [Ca2+]m increases? One suggestion was that, mitochondrial Ca2+ can accumulate directly from the cytosolic Ca2+ pool for prolonged periods, even if the mean [Ca2+]c elevation does not exceed the micromolar level (Szabadkai et al., 2001; Pitter et al., 2002). Alternatively, Ca2+ released from the ER can directly influx into mitochondria at the close contact sites between the ER and the mitochondrial outer membrane. Mitochondrial Ca2+ accumulation through this pathway may have a direct effect on destroying mitochondrial membrane potential and the following release of the apoptotic proteins (Szalai et al., 1999; Hajnoczky et al., 2000; Pacher and Hajnoczky, 2001; Rapizzi et al., 2002; Csordas et al., 2002). It was suggested that a major target of mitochondrial Ca2+ regulation is the mitochondrial permeability transition pore (PTP) (Bernardi and Petronilli, 1996). PTP was thought to be involved in the process of releasing mitochondria-derived apoptotic factors such as cyt c (see review by (Green and Reed, 1998)). Recently, Rizzuto’s group reported that overexpressing VDAC (voltage-dependent anion channel), one of the components of the PTP, could enhance IP3R-mediated mitochondrial Ca2+ uptake and potentiate the apoptotic effect of ceramide (Rapizzi et al., 2002). However, whether the opening of PTP is truly responsible for the release of mitochondrial proteins is highly controversial (Zamzami et al., 1996; Narita et al., 1998; Gao et al., 2001; Zamzami and Kroemer, 2001; Kuwana et al., 2002; De Giorgi et al., 2002; Scorrano et al., 2002). On the basis of the results of our own studies (Gao et al., 2001), we suspect that PTP is more likely to play a role in mitochondrial Ca2+ transport than releasing cyt c. The mechanism of the ER-mitochondria Ca2+ communication is still under active investigation. There is evidence suggesting that mitochondria have buffering effect on cytosolic Ca2+ signals. For instance, in motor neuron cytosolic Ca2+ and mitochondria Ca2+ elevation were observed under electrical stimuli; applying CCCP (carbonyl cyanide m-chlorophenyl hydrazone) which dissipate the proton electrochemical gradient across the mitochondrial membrane can inhibit the increase of [Ca2+]m (David et al., 1998). It was suggested that mitochondrial uptake of Ca2+ contributes to buffering presynaptic cytosolic Ca2+ (David et al., 1998). Another hypothesis called ‘‘hotspot hypothesis’’ showed that the mitochondria Ca2+ uptake was through the close apposition (in the 100 nm range) between ER and mitochondria and in particular between the IP3R and mitochondria (Rizzuto et al., 1998). By using targeted fluorescent proteins and digital deconvolution microscopy, Rizzuto et al. imaged the three-dimensional organization of mitochondria and ER in eukaryotic cells (Rizzuto et al., 1998). They showed evidence of closed localization between ER and mitochondria. They further suggested that the close appositions between ER and mitochondria may represent the site where microdomains of high [Ca2+] are generated upon IP3-mediated Ca2+ release. Mitochondrial Ca2+ uptake may also play an important role on regulating ER Ca2+ refilling after IP3induced ER Ca2+ release. A study by Demaurex’s group showed that in HeLa and HEK 293 cells, some of mitochondria are very close to the sites of Ca2+ release and recycle a substantial portion of the captured Ca2+ back to the vicinal ER domains (Arnaudeau et al., 2001). This hypothesis is consistent with a recent report that the Ca2+ shuttling between mitochondria and ER plays a pacemaker role in the generation of Ca2+ oscillations (Ishii et al., 2006).

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Bcl-2 Family Proteins and Ca2+ Signaling

It is well known that Bcl-2 family proteins play a major role in controlling the permeabilization of the mitochondrial outer membrane during apoptosis. Recently, there was evidence indicating that Bcl-2 family proteins are also involved in modulating the Ca2+ signal in apoptotic cells. First, it was found that some of the Bcl-2 family proteins can localize not only to the mitochondria, but also to other intracellular membranes such as the ER and nuclear envelope. Second, there was indication that some of the Bcl-2 family proteins could interact with the IP3 receptors at the ER. > Table 33-1 summarizes the reported findings on the interplay between the Bcl-2 family proteins and Ca2+ signaling.

4.1 Bcl-2 So far, most of the works on examining the crosstalk between the Bcl-2 family proteins and Ca2+ signal had been carried out on Bcl-2. Starting from a decade ago, several studies have indicated that Bcl-2 may affect ER calcium homeostasis [(Baffy et al., 1993; Magnelli et al., 1994); for review, see (Distelhorst and Shore, 2004)]. However, the reported results were controversial and the mechanisms were not clear. For example, Distelhorst and co-workers reported that Bcl-2 can inhibit Ca2+ release from the ER and thereby preserve the ER calcium pool (Distelhorst and McCormick, 1996; He et al., 1997). This inhibition was not

. Table 33-1 Bcl-2 family proteins and Ca2+ signalling Bcl-2 family proteins Bcl-2

Localization Mitochondria Yes

Calcium signalling ER Yes1,2

Mitochondria ↑ Mitochondria Ca2+ uptake capacity3–5

↓ Secondarily mitochondria Ca2+ uptake6 Bax/Bak

Yes

Yes11,12

↑ Secondarily mitochondria Ca2+ uptake12–14

Bcl-xl

Yes

No2

Unknown

Bik

Yes

Yes19,20

Unknown

Mcl-1

Yes

No

↓ Mitochondria Ca2+ uptake22

ER ↓ Steady-state luminal Ca2+ level6–8 ↓ Agonist-dependent Ca2+ fluxes 6,8,9 ↓Capacitative Ca2+ entry6,7 ↓ Calreticulin expression10 ↓ SERCA2b expression10 ↑ Steady-state luminal Ca2+ level12,15 ↑ Agonist-dependent Ca2+ fluxes12–16 ↓ Type I IP3R expression17 ↓ Agonist-dependent Ca2+ fluxes17,18 Bind to IP3R18 ↑ Agonist-dependent Ca2+ fluxes21 No effect22

References in the table: 1 Chen-Levy et al. (1989); 2Kaufmann et al. (2003); 3Murphy et al. (1996); 4Ichimiya et al. (1998); 5Zhu et al. (1999); 6Pinton et al. (2000); 7Foyouzi-Youssefi et al. (2000); 8Palmer et al. (2004); 9Chen et al. (2004); 10Vanden Abeele et al. (2002); 11Zong et al. (2003); 12Scorrano et al. (2003); 13Nutt et al. (2002a); 14Chami et al. (2004); 15Oakes et al. (2005); 16Nutt et al. (2002b); 17Li et al. (2002); 18White et al. (2005); 19Germain et al. (2002); 20Germain et al. (2005); 21Mathai et al. (2005); 22Minagawa et al. (2005)

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due to an alteration in IP3Rs expression or luminal Ca2+ concentration, but was mediated through a functional interaction between Bcl-2 and IP3Rs that inhibits their channel opening in response to IP3 activation (Chen et al., 2004). In contrast, other groups reported that Bcl-2 overexpression caused a reduction of the steady-state. Ca2+ concentration in the ER, and thus the amount of agonist-releasable Ca2+ flux was less than the control. This reduction was mainly due to an increase in the Ca2+ leak across the ER membrane (FoyouziYoussefi et al., 2000; Pinton et al., 2000; Palmer et al., 2004) (These two alternative hypotheses are summarized in > Figure 33-6). Recently, it became technically possible to directly measure ER luminal

. Figure 33-6 Regulation of Ca2+ release from ER by Bcl-2. Two possible ways in explaining the effect of Bcl-2 on ER Ca2+ signaling had been suggested: (A) Reduce traffic. Bcl-2 was shown to reduce the ER Ca2+ release because of a functional interaction of Bcl-2 with IP3Rs that inhibited their channel opening in response to IP3. (B) Reduce source. It was suggested that Bcl-2 can decrease the steady-state Ca2+ concentration in the endoplasmic reticulum (ER) by increasing the Ca2+ leakage across the ER membrane. It is not yet clear whether the enhanced leakage of Ca2+ is through channels formed by Bcl-2 itself or through the interaction of Bcl-2 with existing channels, such as IP3R or RyR. (Reproduced from Guo et al., 2005)

calcium concentration. It was reported that manipulations leading to decreased [Ca2+]ER often protected cells against apoptosis, whereas manipulations leading to increased [Ca2+]ER often sensitized cells to apoptosis (Nakamura et al., 2000; Lilliehook et al., 2002). In addition, it was reported that Bcl-2 can increase the capacity of mitochondria in neurons to store Ca2+, presumably by preventing the opening of the PTP (Murphy et al., 1996). Thus, by localizing to both the ER and mitochondria, Bcl-2 may prevent Ca2+ mobilization between the two organelles by three possible mechanisms: (1) lowering the pool of free Ca2+ in the ER, (2) inhibiting Ca2+ release from the ER through interaction with IP3Rs, or (3) increasing the tolerance of mitochondria to high Ca2+ loads.

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4.2 Bax/Bak It was known that pro-apoptotic Bax not only interact with mitochondria but also with ER. For example, it was reported that Bax can localize to both ER and mitochondria in PC-3 prostate cancer cells (Nutt et al., 2002a). Bax-mediated alterations in ER and mitochondrial Ca2+ levels were regarded as important upstream signals for cytochrome c release in some apoptotic systems (Nutt et al., 2002b). ER stress induced by thapsigargin or tunicamycin can induce Bax conformational change and oligomerization on ER, followed by caspase-12 activation (Zong et al., 2003). Korsmeyer and co-workers reported that Bax/Bak double-knockout (DKO) cells had a reduced resting ER calcium level, and resulted in decreasing Ca2+ mobilization from ER to mitochondria (Scorrano et al., 2003). They proposed that the reduced ER calcium is due to the increased ER calcium leak by hyperphosphorylated IP3R-1. Knocking down Bcl-2 somehow could counter act this effect; it decreased IP3R-1 phosphorylation and ER calcium leakage rate in the Bax/ Bak DKO cells (Oakes et al., 2005). Recently, we have examined the effects of Bax on ER Ca2+ release in HeLa cells. Our results suggested that Bax could interact with IP3 receptors to enhance the Ca2+ efflux from ER under apoptotic stress (unpublished results).

4.3 Other Bcl-2 Family Proteins Other Bcl-2 family proteins such as Bcl-xL, Bik were also reported to affect Ca2+ signaling. Bcl-xL is an antiapoptotic protein. Similar to Bcl-2, Bcl-xL could prevent apoptosis initiation by antagonizing the oligomerization of Bax/Bak (Minn et al., 1999; Tan et al., 1999). Bcl-xL was found to interact with the carboxyl terminus of the IP3R and sensitize single IP3R channel in ER membrane to low [IP3]. As a result, Bcl-xL reduced [Ca2+]ER. The pro-apoptotic proteins Bax and tBid could antagonize this effect by blocking the biochemical interaction of Bcl-xL with IP3R (White et al., 2005). Thus, these results suggested that Bcl-xL is a direct effector of the IP3R, increasing its sensitivity to IP3 and enabling ER Ca2+ release to be more sensitively coupled to extracellular signals. Bik (Bcl-2 interacting killer) is a pro-apoptotic BH3-only Bcl-2 family protein (Elangovan and Chinnadurai, 1997). Functional study showed that Bik induced cell death through the BH3 domain by forming heterodimerization. Gordon and co-workers showed that Bik can interact with Ca2+ signaling (Germain et al., 2002, 2005). First, they found that ER localized-Bik could trigger Bax translocate to mitochondria followed by cyt c release. Second, they showed that Bik can induce ER Ca2+ release which is dependent on Bax and Bak. Mcl-1 (myeloid cell leukaemia-1) is an anti-apoptotic protein which was originally identified in 1993 in differentiating myeloid cells (Kozopas et al., 1993). It was reported that Mcl-1 could interact with pro-apoptotic proteins, such as Bax, Bak, and Bad, and neutralize their effects (Gelinas and White, 2005). Expression of Mcl-1 did not affect the level of ER Ca2+ store or the expression level of IP3Rs. However, [Ca2+]m induced by apoptotic stimuli were decreased in cells overexpressing Mcl-1 (Minagawa et al., 2005). These findings suggested that Mcl-1 directly inhibits Ca2+ signals within mitochondria and thus may provide a novel mechanism for preventing apoptosis.

5

Ca2+ Signaling and Neuronal Cell Death

5.1 Ca2+ and ROS-Induced Neuronal Cell Death Neuronal cell death is a common feature of many diseases, including brain ischemia, neuronal degenerative diseases and traumatic injury. Thus, it is important to investigate the mechanisms that mediate neuronal cell death. A number of factors are known to contribute to brain disorders. Among them, oxidative damage and Ca2+ dysregulation play particularly important roles in the pathogenesis of these diseases (Chinopoulos and Adam-Vizi, 2006; Mattson, 2006, 2007).

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Oxidative stress is the excessive oxidation of cellular biomolecules that leads to cellular damage, which is caused by reactive oxygen species (ROS) including superoxide radicals, hydrogen peroxide, hydroxyl radicals, nitric oxide, and metabolites thereof (Nordberg and Arner, 2001). The brain, which consumes large amount of oxygen, is vulnerable to oxidative damage. ROS can oxidize critical cellular components such as membrane lipids (Yla-Herttuala, 1999), proteins (Stadtman and Levine, 2000), and DNA (Marnett, 2000); these, lead to apoptosis or necrosis. In addition, the toxic by-products generated during oxidative damage also contribute to brain injury. There is increasing evidence for the association between ROS production, the induction of apoptosis (or necrosis) and the pathogenesis of stroke and neurodegenerative disorders (Loh et al., 2006). Redox modulation of Ca2+-regulating proteins can occur in different physiological and pathological processes, resulting in altered amplitude and spatiotemporal characteristics of Ca2+ signals. Exogenous ROS could induce dynamic changes in [Ca2+]c in a variety of types of cells (Doan et al., 1994; Kumasaka et al., 1999; Suzuki and Ford, 1999; Gen et al., 2001). This effect could be due to mobilization of Ca2+ from intracellular Ca2+ stores or by altering influx of extracellular Ca2+. Functionally, the effect of ROS on Ca2+ signaling can vary from stimulatory to inhibitory, depending on the type of target proteins, the type of oxidants, the dose, duration of exposure, and the cell conditions. The regulation of intracellular Ca2+ distribution and fluxes by ROS is very complex (> Figure 33-7). Furthermore, excess intracellular Ca2+ can lead to generation of free radicals within vulnerable neuronal populations which in turn induces Ca2+ dyshomeostasis (Mattson et al., 1995). In this section, we briefly discuss how ROS modify key Ca2+ signaling proteins and how the resultant intracellular Ca2+ signaling changes may contribute to neuronal cell death. . Figure 33-7 A diagram summarizing the possible mechanisms of ROS-induced elevation of [Ca2+]c. ROS could alter the cytosolic Ca2+ level by affecting a number of Ca2+-regulators associated with different subcellular structures, including the ER and plasma membrane. These Ca2+-regulating proteins include various Ca2+ channels (such as IP3R, VDCC and TRPM2 channels) and Ca2+ transporters (such as Ca2+-ATPases and Na+/Ca2+ exchangers). Also, ROS can cause overloading of Ca2+ in the mitochondria, which could result in more free radical production. (Based on Guo et al., 2005)

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5.2 ROS-Regulated Plasma Membrane Ca2+ Signaling Pathways in Neuronal Cell Death Eukaryotic cells can import Ca2+ through a number of gated plasma membrane channels. These Ca2+ signals transmit information to a large number of targets (e.g., enzyme). Later, these excessive Ca2+ ions must be exported again by certain membrane transport proteins. Many of these plasma membrane Ca2+ regulators can be altered during ROS-induced cell death. TRPM. Members of the melastatin subfamily of TRP proteins (TRPM), particularly TRPM2, may play key roles in neuronal death activated by oxidative stress (Waring, 2005). It was reported that TRPM2, a long TRP channel containing an ADP-ribosylase domain, could be activated by H2O2 and agents that produce reactive oxygen/nitrogen species (Hara et al., 2002; Wehage et al., 2002). However, whether it was the direct or indirect effect of ROS is still debated. Recent work on TRPM2 suggested a role for ADP-ribose, which is produced after poly(ADP-ribose) polymerase (PARP) activation, as a mediator between oxidative damage and downstream TRPM2 activation (Fonfria et al., 2004). Application of PARP inhibitors was shown to reduce ADPR formation and inhibit TRPM2 activation as well as Ca2+ entry (Miller, 2004). Furthermore, suppressing TRPM2 using inhibitors or knocking down TRPM2 with anti-sense RNAs could protect cells significantly against death compared with nontreated control cells. These results indicated that H2O2 may cause cell death through a TRPM2-dependent mechanism (Hara et al., 2002; Zhang et al., 2003, 2006; Fonfria et al., 2005; Kaneko et al., 2006). Thus, elucidation of the signaling properties of TRPM family members may provide useful information in unmasking the true nature of neuronal death following CNS (central nerve system) injury. VDCC (voltage-dependent Ca2+ channel). Many studies have suggested that ROS can modulate the activity of VDCCs. For example, oxidation by external H2O2 of the a1A, a2/Dd, and b3 subunits of P/Q type VDCC channels heterologously expressed in Xenopus oocytes could enhance the inward whole-cell Ca2+ currents carried by this VDCCs subtype (Li et al., 1998). Furthermore, the lipid peroxidation product hydroxynonenal (HNE) caused opening of L-type channels, increased intracellular Ca2+ concentration and neuronal cell death. This effect could be abrogated by applying the L-type channel blocker, nimodipine (Lu et al., 2002). There was indirect evidence that VDCCs could play an important role in oxidative stress-induced neuronal cell death. For example, L-type channels were shown to be involved in the increased calcium fluxes caused by amyloid-b peptides and the cell death so induced could be abrogated by radical scavengers (Green and Peers, 2001). In addition, in the secretory lipases sPLA2-IIA (secretory phospholipase A2-IIA) induced cell death in neurons, it appeared that ROS production induced by the lipase activity could activate L-type channels and result in Ca2+ overloading in the neuron (Yagami et al., 2003). PMCA. Plasma membrane Ca2+ ATPases (PMCA) can be inhibited by oxidants (Ermak and Davies, 2002). For example, exposure of red blood cell membranes to ferrous sulfate, which generates hydrogen peroxide and hydroxyl radicals, resulted in a concentration- and time-dependent inhibition of the PMCA (Rohn et al., 1993). This inhibition can be prevented by certain free-radical scavengers. In addition, studies in rat brain synaptosomes showed that micromolar level of peroxynitrite inhibit 75% of the activity of PMCA within 50 min, the inhibition is accompanied by parallel increase of calcium level in the vesicles (Gutierrez-Martin et al., 2002). It was also found that PMCA purified from red blood cell membranes was inactivated by hydrogen peroxide because of cross-linking of thiol residues (Zaidi et al., 2003). It was suggested by Kip et al. that PMCA may play an important role in Ca2+-mediated neuronal cell death. They studied the short-term effect of oxidative stress on the plasma membrane Ca2+ extrusion systems in hippocampal neurons and found that after 2–3 h exposure to 300 mM H2O2, all PMCAs were significantly downregulated at the mRNA and protein levels. Rapid internalization and aggregation of the PMCA was also observed. This would reduce the capacity for Ca2+ extrusion. This finding suggested that the plasma membrane calcium systems are sensitive early targets of neurotoxic oxidative stress (Kip and Strehler, 2007).

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5.3 ROS-Regulated ER Ca2+ Signaling Pathways in Neuronal Cell Death SERCA. Redox agents are known to modulate SERCA (Sarco-endoplasmic reticulum Ca2+ ATPase). In contrast to the IP3R, the activity of SERCA was inhibited by oxidants, whereas reducing agents (e.g., DTT and GSH) protected SERCA from this inhibition (Zima and Blatter, 2006). The studies on the mechanism of how oxidants regulate the activity of SERCA were mainly in sarcoplasmic reticulum (SR) or microsomes. It was shown that O2 and H2O2/OH inhibited Ca2+ uptake into the SR (Rowe et al., 1983; Kukreja et al., 1988; Castilho et al., 1996; Morris and Sulakhe, 1997; Xu et al., 1997). It seems that SERCA can be inhibited both by oxidation of its sulfhydryl groups (Scherer and Deamer, 1986; Suzuki and Ford, 1991; Morris and Sulakhe, 1997) and by direct attack of oxidants on the ATP binding site (Xu et al., 1997). Furthermore, ROS could cause inhibition of SERCA function by uncoupling Ca2+ uptake activity from ATP hydrolysis (Rowe et al., 1983). Additionally, ROS could inhibit the activity of membrane-bound enzymes by peroxidation of membrane phospholipids (Morris and Sulakhe, 1997). Since ROS is involved in many pathological clinical situations of brain disorders, such as ischemia-reperfusion injury, ROS-induced downregulation of SERCA plays a significant role in neuronal cell death (Larsen et al., 2005). IP3 receptors. It had been shown that redox status could modulate IP3R activities in various cell types. The IP3R can be modulated by ROS directly. For example, in vascular smooth muscle cells, superoxide could sensitize Ca2+ release stimulated by IP3 (Suzuki and Ford, 1992). Oxidized glutathione (GSSG) (Henschke and Elliott, 1995) and H2O2 (Wesson and Elliott, 1995) have also been shown to induce Ca2+ release from IP3-mediated stores in endothelial cells. Additionally, in permeabilized hepatocytes, GSSG stimulated the IP3R by increasing the binding affinity of IP3 to the IP3R (Renard-Rooney et al., 1995). In an in vivo study of the effect of hypoxia on IP3 receptors in piglets, it was shown that IP3 binding to receptor increased in hypoxic brain and this was inhibited in animals treated with N-nitro-L-arginine, an inhibitor of nitric oxide synthase (Mishra et al., 2003). Such an effect was considered to be a major contributing factor in subsequent calcium-dependent neuronal apoptotic cell death. It was also demonstrated that the release of Ca2+ from the ER, mediated by both RyR and IP3R, was involved in amyloid-b toxicity and could contribute, together with the activation of other intracellular neurotoxic mechanisms, to the amyloid-b-induced neuronal death (Ferreiro et al., 2004, 2006).

6

Conclusion and Perspectives

From this review, it is clear that Ca2+ signal plays a very important role in the apoptotic process. Particularly, we have convincing evidence that Ca2+ signal is involved in regulating the initiation phase of apoptosis, acting upstream of the dysfunction of mitochondria. At present, the exact downstream target of the Ca2+ signal is not yet clear. It appears that there are multiple targets for the Ca2+ signal, including various types of calpains and calcineurin. Furthermore, excessive Ca2+ uploading in the mitochondria matrix can directly damage the mitochondria and thus results in releasing apoptotic factors such as cyt c. We think that the targets of the Ca2+ signal probably depends on the cell model and the specific type of apoptotic stress. One of the most exciting findings in recent years in this field is the discovery that Ca2+ flux released from the ER can be modulated by the Bcl-2 family proteins. Previously, the major role of Bcl-2 family proteins in apoptosis was thought to be regulating the permeabilization of the mitochondrial outer membrane. The newly discovered interaction between the Bcl-2 family proteins and the Ca2+ regulators offers a second signaling mechanism in controlling cell death. This discovery also further demonstrates the importance of Ca2+ signaling in apoptosis. Finally, there is strong evidence indicating that ROS-induced neuronal cell death is dependent on elevation of free Ca2+ in the cytosol. Part of the Ca2+ flux can be released from internal stores such as ER, while part of the Ca2+ elevation can be caused by inward Ca2+ flux passing through the plasma membrane. So far, a number of Ca2+-regulating proteins responsible for controlling this external Ca2+ flux has been identified. It would be interesting to investigate which of these Ca2+ regulators play the major role in

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modifying the Ca2+ homeostasis in a particular pathological disorder. By finding specific inhibitors for the targeted Ca2+ regulator, such research may enable us to devise new drug treatments for some major neurodegenerative diseases.

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