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ISBN: 978-0-12-374805-8

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CONTRIBUTORS

Zakaria A. Almsherqi Department of Physiology, Yong Loo Lin School of Medicine, National University of Singapore, 117597 Singapore Angelika Barnekow Department of Experimental Tumorbiology, University of Mu¨nster, Badestraße 9, 48149 Mu¨nster, Germany Yuru Deng Department of Physiology, Yong Loo Lin School of Medicine, National University of Singapore, 117597 Singapore Robbin D. Eppinga Department of Biology, University of Iowa, Iowa City, Iowa 52242 Andrea C. Gore Institute for Cellular and Molecular Biology, and Division of Pharmacology and Toxicology, and Institute for Neuroscience, University of Texas, Austin, Texas 78712 Jian-Ping Jin Section of Molecular Cardiology, Evanston Northwestern Healthcare and Northwestern University Fienberg School of Medicine, Evanston, Illinois 60201 Daniel Kessler Department of Experimental Tumorbiology, University of Mu¨nster, Badestraße 9, 48149 Mu¨nster, Germany Sepp D. Kohlwein Institute of Molecular Biosciences, University of Graz, A8010 Graz, Austria Tomas Landh Novo Nordisk A/S, DK-2760 Ma˚løv, Denmark Yan Li Department of Biology, University of Iowa, Iowa City, Iowa 52242 Jim Jung-Ching Lin Department of Biology, University of Iowa, Iowa City, Iowa 52242 Jacqueline A. Maffucci Institute for Neuroscience, University of Texas, Austin, Texas 78712 vii

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Contributors

Fernando Martı´n-Belmonte Centro de Biologı´a Molecular Severo Ochoa, Consejo Superior de Investigaciones Cientı´ficas-UAM, Madrid 28049, Spain Alejo E. Rodrı´guez-Fraticelli Centro de Biologı´a Molecular Severo Ochoa, Consejo Superior de Investigaciones Cientı´ficas-UAM, Madrid 28049, Spain Chieko Saito Molecular Membrane Biology Laboratory, Advanced Science Institute, RIKEN, Saitama 351-0198, Japan Anika Thyrock Department of Experimental Tumorbiology, University of Mu¨nster, Badestraße 9, 48149 Mu¨nster, Germany Takashi Ueda Laboratory of Developmental Cell Biology, Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo 113-0033, Japan Qinchuan Wang Department of Biology, University of Iowa, Iowa City, Iowa 52242

C H A P T E R

O N E

Roles of Caldesmon in Cell Motility and Actin Cytoskeleton Remodeling Jim Jung-Ching Lin,* Yan Li,* Robbin D. Eppinga,* Qinchuan Wang,* and Jian-Ping Jin† Contents 1. Introduction 2. Organization and Regulation of Caldesmon (CALD1) Gene 2.1. Caldesmon gene organization 2.2. Regulation of CALD1 gene expression 3. Structure and Function Study of CaD 3.1. Domains for the bindings to CaD-interacting proteins 3.2. Phosphorylated residues on CaD by various kinases 3.3. CaD and calponin interaction 4. CaD and Regulation of Smooth Muscle Contraction 5. CaD and Regulation of Nonmuscle Cell Motility and Cytoskeleton Dynamics 5.1. Intracellular localization of l-CaD and its functions 5.2. Regulation of l-CaD activity in nonmuscle cells 6. CaD, Podosomes, and Diseases 7. Concluding Remarks Acknowledgments References

2 3 3 4 7 8 14 15 24 27 28 30 41 44 51 51

Abstract Caldesmon (CaD) is a multimodular protein that regulates contractility and actin cytoskeleton remodeling in smooth muscle and nonmuscle cells. A single gene (CALD1) encodes high molecular mass CaD (h-CaD) and low molecular mass CaD (l-CaD) by alternative splicings. The h-CaD exclusively expresses in smooth muscle, whereas the l-CaD ubiquitously expresses in all cell types except skeletal muscle. The h-CaD/l-CaD ratio could be a marker for monitoring differentiating and pathological states of smooth muscles. The l-CaD associates with stress fibers

* {

Department of Biology, University of Iowa, Iowa City, Iowa 52242 Section of Molecular Cardiology, Evanston Northwestern Healthcare and Northwestern University Fienberg School of Medicine, Evanston, Illinois 60201

International Review of Cell and Molecular Biology, Volume 274 ISSN 1937-6448, DOI: 10.1016/S1937-6448(08)02001-7

#

2009 Elsevier Inc. All rights reserved.

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and membrane ruffles in nonmuscle cells and with the actin core of podosomes in highly motile/invasive cells. Together with tropomyosin, CaD stabilizes actin filaments and inhibits actin-tropomyosin-activated myosin ATPase activity. This inhibition can be effectively released by Ca2þ–calmodulin and/or by phosphorylation with various kinases. Through its interactions with a spectrum of actin-binding proteins, CaD modulates dynamics of cortical actin networks and stress fibers, which are essential to cell motility and cytoskeleton rearrangement. Regulation of CaD level and its activity may provide a novel strategy for gene therapy.

1. Introduction Caldesmon (CaD) is a myosin-, actin-, tropomyosin-, and Ca2þ– calmodulin-binding protein, capable of regulating actomyosin contraction, actin filament dynamics and cytoskeleton remodeling in smooth muscle and nonmuscle cells. It was first discovered from chicken gizzard as an actin- and Ca2þ–calmodulin-binding protein (Sobue et al., 1981). Subsequently, it was found to be ubiquitous in thin filaments of smooth muscles and in actin microfilaments of nonmuscle cells. Association of CaD to tropomyosincontaining actin filaments effectively inhibits the actomyosin ATPase activity and the in vitro actin filament motility. This inhibition can be reversed by Ca2þ–calmodulin and/or phosphorylation by various kinases, including casein kinase II (CKII), protein kinase C (PKC), Ca2þ–calmodulin kinase II (CaMKII), extracellular signal-regulated kinase (ERK), p21-activated kinase (PAK), oncogenic v-erbB tyrosine kinase, and Cdc2 kinase. Biochemical and physiological evidence to support this general conclusion has been previously reviewed in many excellent review articles (Bretscher, 1986; Chalovich, 1988; Chalovich et al., 1998; Dabrowska et al., 2004; Hai and Gu, 2006; Huber, 1997; Kakiuchi and Sobue, 1983; Marston and Redwood, 1991; Matsumura and Yamashiro, 1993; Morgan and Gangopadhyay, 2001; Sellers, 1999; Sobue and Sellers, 1991; Sobue et al., 1988; Wang, 2001; Yamashiro and Matsumura, 1991; Yamashiro et al., 1994). In this review, we will briefly summarize what we know about caldesmon gene organization and expression regulation, functional domains and phosphorylation sites, as well as effects of CaD on other cytoskeletal proteins and phospholipids. We will then discuss roles of CaD in smooth muscle contraction upon agonist stimulation, in actin cytoskeleton remodeling during nonmuscle cell migration and cytokinesis, as well as in podosome dynamics and formation. Regulation of CaD activity is important in controlling actomyosin ATPase activity and actin cytoskeleton remodeling for normal cell function and in certain diseases, such as transformed cell proliferation and motility, brain tumors (e.g., gliomas), bladder outlet obstruction, glaucoma, angiogenesis, and endothelial cell barrier dysfunction.

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2. Organization and Regulation of Caldesmon (CALD1) Gene 2.1. Caldesmon gene organization Two classes of CaD proteins, h-CaD (high molecular mass CaD) and l-CaD (low molecular mass CaD), are preferentially expressed in smooth muscle and nonmuscle cells, respectively. Alternative splicing of a single Caldesmon (CALD1) gene generates l-CaD isoforms, which lack a highly repetitive central helical domain found in h-CaD (Haruna et al., 1993; Hayashi et al., 1992; Payne et al., 1995). In addition, two distinct promoters are used in different cell types or tissues to generate l-CaD isoforms with distinct N-termini (Hayashi et al., 1992; Humphrey et al., 1992, 1995; Novy et al., 1991; Payne et al., 1995; Yano et al., 1994, 1995). In humans, the CALD1 gene is located on chromosome 7q33 (Hs.490203, http://www. ncbi.nlm.nih.gov/UniGene/clust.cgi) and consists of 17 exons (Fig. 1.1). Through the use of alternatively spliced exons (exons 3a, 3b, and 4) and distinct promoters (starting with exon 1a-1, -2, -3, or exon 1b), it gives rise to aorta h-CaD (793 amino acid (aa) residues, 93.2 kDa), WI-38 l-CaD I (564 aa, 65.7 kDa), WI-38 l-CaD II (538 aa, 62.7 kDa), HeLa l-CaD I (558 aa, 64.3 kDa), and HeLa l-CaD II (532 aa, 61.2 kDa) (Hayashi et al., 1992; Humphrey et al., 1992; Novy et al., 1991). The exon 3 contains an intraexonic 50 splicing site and generates exons 3a and 3b. All l-CaD isoforms either from WI-38 fibroblasts or HeLa cells do not use exon 3b (Fig. 1.1). At the present time, the significance of two l-CaD isoforms in the same cell remains to be determined. Similar genomic structure and isoform expression are also observed in chicken and other mammalian Cald1 genes (Haruna et al., 1993; Payne et al., 1995; Yano et al., 1994, 1995). In chicken Cald1, there is an additional exon (34 bp) present between the last two exons. Also there is an additional alternative 30 -splice site at the intron/exon junction of the last exon, which generates different l-CaD isoforms in chicken gizzard and brain (Haruna et al., 1993; Sobue et al., 1999). The full-length cDNA sequences obtained from chicken gizzard h-CaD by Hayashi et al. (1989) and Bryan et al. (1989) were later proven to be identical, each encoding a 771-amino acid (88.7 kDa) protein (Guo et al., 1999). Thus, the residue numbers for some of functional domains and phosphorylation sites on chicken CaD isoforms used here reflect this finding of Guo et al. (1999), which have a 15-residues difference from that reported in much of the earlier literature. Due to a high content of glutamine residues, h-CaD and l-CaD isoforms migrate anomalously slowly during SDS-PAGE and have apparent molecular masses of 120–130 kDa and 70–80 kDa, respectively (Graceffa et al., 1988).

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Human caldesmon gene

1a-1

1a-2

1a-3

ATG

1b

2

3a3b

4

5

ATG

6

7

89 1011

12

13

TGA polyA

Aorta h-CaD WI-38 I-CaD I WI-38 I-CaD II HeLa I-CaD I HeLa I-CaD II

Figure 1.1 Schematic diagram of the human caldesmon (CADL1) gene organization and isoform diversity. The 17 exons are indicated by the colored boxes, the introns in between are presented by solid lines, and the spliced out intron are represented by the dashed lines. The isoforms that are listed under the gene have been cloned from human aorta, HeLa cells and WI-38 fibroblast cells. The low molecular mass CaD (l-CaD) isoforms use two different promoters to produce different N-terminus of l-CaD from exon 1a-3 or 1b (also called exon 10 or 1, respectively, in Payne et al. (1995); and Zheng et al. (2004); however, none of l-CaD contains exon 3b. The high molecular mass CaD (h-CaD) uses all exons except exon 1b. A full-length cDNA for h-CaD isoform has also been cloned from HeLa cells (Cuomo et al., 2005), however, its amino acid sequence is identical to aorta smooth muscle h-CaD (Hayashi et al., 1992).

2.2. Regulation of CALD1 gene expression Decreased expression of l-CaD at both message and protein levels has been found in many transformed and cancer cells (Novy et al., 1991; Owada et al., 1984; Ross et al., 2000; Tanaka et al., 1993). Although the significance of this reduced expression is not completely understood, it may account for the loss of stress fibers and focal adhesions observed in these cells and possibly for facilitating their morphological changes and motility. The recent findings that CaD is an integral component of podosomes (sites of cell adhesion and active extracellular matrix remodeling) and that CaD is capable of modulating podosome assembly and dynamics (Eves et al., 2006; Gu et al., 2007; Morita et al., 2007) further support this concept. Transformed cells generally lose their relatively immobile focal adhesions and often develop into more dynamic podosomes to facilitate invasion. We will further discuss the advances in understanding CaD’s role in regulating podosomes formation in Section 6. The decrease of l-CaD protein has been also observed in differentiating myoblast L6 cells until no detection of l-CaD in the

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differentiated myotubes (Yamashiro-Matsumura et al., 1988). In addition to the regulation of CaD expression during myogenesis, chicken myotubes are shown to functionally exclude microinjected l-CaD proteins (Yamakita et al., 1990). Similarly, differentiated smooth muscle cells express exclusively h-CaD, whereas dedifferentiated smooth muscle cells associated with some pathological conditions such as atherosclerosis, hypertension synthesize a decreasing amount of h-CaD, concomitant with an increasing expression of l-CaD (Ueki et al., 1987). The decrease of l-CaD in transformed and differentiated cells may be a direct result of decreased Cald1 gene transcription and/or alternative splicing, in addition to a possible reduction in mRNA stability. We will discuss what we know about the regulations of alternative splicing and promoter activity in the following sections. 2.2.1. Regulation of 50 -splice site selection in exon 3 of human CALD1 gene As shown in Fig. 1.1, all of l-CaD isoforms lack exon 3b as a result of the usage of an intraexonic 50 -splice site in exon 3. The selection of this intraexonic 50 -splice site or its downstream (terminal) 50 -splice sites would determine the expression of l-CaD or h-CaD, respectively, isoform. Studies with human CALD1 gene, Humphrey et al. have shown that the usage of the intraexonic 50 -splice site requires an exon-splicing enhancer (a long purine-rich sequence) within exon 3b, which consists of four nearly identical 32-nucleotide repeats (Humphrey et al., 1995). In fact, only one purinerich repeat is sufficient to promote this intraexonic splice site selection and to repress the choice of the terminal splice site. When this purine-rich enhancer sequence (called exon-splicing enhancer) is placed downstream of the terminal 50 -splice site, it has no effect on splice site choice. The exonsplicing enhancer in human CALD1 gene has high sequence homology to a general consensus sequence, PRESE (purine-rich exon-splicing enhancer), found in many alternative splicing genes to regulate the splice site selection (Elrick et al., 1998; Kakizuka et al., 1990; Lavigueur et al., 1993; Ryner et al., 1996; Tanaka et al., 1994; Watakabe et al., 1993). Similarly, there are nine closely conserved PRESE repeats found in the exon 3b of chicken Cald1 gene, which can also serve as an exon-splicing enhancer to determine splice site selection. Although it remains to be determined, the regulatory factor(s) may exist in nonmuscle cells to recognize this exon-splicing enhancer and promote the expression of l-CaD isoform. On the other hand, in differentiated smooth muscle cells, lack of these putative regulatory factors would result in the expression of the h-CaD isoform. 2.2.2. Transcriptional regulation of chicken Cald1 gene Two types of promoters have been detected in chicken Cald1 gene. The gizzard-type promoter is located upstream of exon 1a-1 and is used in the expression of gizzard and fibroblast CaDs. The brain-type promoter locates

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at the upstream region of exon 1b, and is used in the expression of brain l-CaD (Yano et al., 1994, 1995). In vitro promoter activity tested in chicken embryo fibroblasts reveals that the gizzard-type promoter has about 10-fold higher activity than the brain-type promoter. Furthermore, a single CArG box located between 309 bp and 300 bp of the gizzard-type promoter is identified as an essential cis-element for cell type-specific expression of the Cald1 gene (Yano et al., 1995). The functionally important CArG boxes have also been identified in the promoters/enhancers of many smooth muscle-restricted genes, including smooth muscle myosin heavy chain and a-actin, SM22a, h1 calponin, b-tropomyosin, and telokin genes (Miano, 2003; Parmacek, 2001). Serum response factor (SRF), a widely expressed transcription factor, is known to directly bind the consensus CArG box, and initiates the transcription of many immediate early genes for cell growth and many muscle-specific genes for muscle differentiation (Miano, 2003). The molecular mechanism underlying how SRF orchestrates these seemly contradictory gene expression programs remains unclear. However, the discovery that smooth muscle-restricted transcription factors such as Nkx3.2 and GATA-6 are able to form complexes with SRF suggests a novel mechanism to activate smooth muscle genes such as caldesmon, SM22a, and a1-integrin during differentiation (Nishida et al., 2002). In fact, chicken Cald1 promoter contains a GATA-6 binding site (GATA box, at 314 bp), the CArG box (at 309 bp), and a Nkx3.2 binding site (at 544 bp) (Yano et al., 1995). In situ hybridization studies reveal that GATA-6, SRF, and Nkx3.2 transcripts are coexpressed in the medial smooth muscle cell layer, which also exclusively expresses h-CaD. On the other hand, only SRF and Nkx3.2 are found in the adventitial and intimal (both are nonsmooth muscle cells) layers of artery, which do not express h-CaD (Nishida et al., 2002). It appears that a combination of these three transcription factors activate the expression of h-CaD and other smooth muscle genes in vascular smooth muscle cells in a way very similar to that SRF, Nkx2.5, and GATA-4 trans-activate cardiac atrial natriuretic factor (ANF) gene (Durocher and Nemer, 1998). Interestingly, in the visceral smooth muscle such as gizzard, GATA-6 is only expressed in the glandular (nonsmooth muscle) layer and is absent in the smooth muscle cell layer, despite that both SRF and Nkx3.2 are expressed in the gizzard (Nishida et al., 2002). Therefore, different mechanisms or different coregulators must be involved in the expression of CaD in vasculature and visceral smooth muscles. It has been reported that SRF expression is essential for the development of mesoderm-derived muscles, that is, cardiac and smooth muscles (Arsenian et al., 1998). However, SRF is also ubiquitously distributed in nonmuscle cells and is involved in the transcription of a wide variety of genes. These observations suggest that coregulators for SRF would need to confer celltype specific transcription. Recently discovered myocardin is capable of complexing with SRF and provides a novel mechanism for SRF to activate

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smooth muscle genes, including CALD1 (Pipes et al., 2006). Myocardin is a remarkably potent transcriptional coactivator expressed exclusively in cardiac and smooth muscle cells (Du et al., 2003; Parmacek, 2007; Pipes et al., 2006; Wang et al., 2001, 2002). Myocardin by itself does not bind to DNA but contains a transcriptional activation domain. Ectopic expression of myocardin in nonmuscle cells induces smooth muscle differentiation, whereas the loss of myocardin shows embryonic lethality and no evidence for vascular smooth muscle differentiation (Li et al., 2003a). It is further shown that myocardin and phospho-Elk-1 compete for the interaction with a common binding site on SRF, providing a plausible mechanism for SRF activating cell type-specific gene expression (Miralles et al., 2003; Wang et al., 2004; Zhou et al., 2005). Elk-1 is a transcription factor for inducing gene expression in response to serum and growth factors, and is activated upon phosphorylation by mitogen activated protein kinase (MAPK) and stress-activated protein kinase/Jun N-terminal kinase (SAPK/JNK). The mutually exclusive association of myocardin and phospho-Elk-1 with SRF provides a binary switch (a phenotypic modulation) in which growth signals can modulate the proliferative and contractile phenotypes of smooth muscle cells. Differentiated smooth muscle cells do not proliferate, morphologically assemble more contractile apparatus (contractile phenotype), and biochemically express exclusively h-CaD, as well as other smooth muscle markers (Sobue et al., 1999). Both the IGF-1-stimulated phosphoinositide 3-kinase (PI3K) and protein kinase B (PKB)/Akt pathways are known to play a critical role in maintaining the differentiated phenotype of smooth muscle cells (Hayashi et al., 1998). On the other hand, growth factors, such as PDGF, bFGF, and EGF as well as cell stress can activate ERK, p38 MAPK, and SAPK pathways and induce the dedifferentiation (proliferative phenotype) of smooth muscle cells, leading to down-regulation of h-CaD and up-regulation of l-CaD (Hayashi et al., 1999, 2001). Phenotypic modulation of smooth muscle cells is known to play a critical role in atherosclerosis, hypertension and leiomyogenic tumorigenesis (Sobue et al., 1999).

3. Structure and Function Study of CaD CaD is capable of switching thin filaments between ‘‘on’’ and ‘‘off ’’ states, in much the same way as troponin does in striated muscle (Marston et al., 1994a). However, structural studies using electron microscopy and 3D image reconstruction reveal that CaD together with tropomyosin flanks strong sites of myosin interaction on actin filaments without covering them. These interactions are unlike those of troponin–tropomyosin and therefore the mechanism of actomyosin ATPase inhibition by CaD-tropomyosin

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may differ from the by troponin–tropomyosin (Hodgkinson et al., 1997; Lehman et al., 1997). This CaD location on thin filaments or actin microfilaments also suggests that it may be capable of competing or cooperating with a number of actin-binding proteins in the cell to modulate the actin cytoskeleton in response to stimuli (Greenberg et al., 2008). In particular, evidence has shown that Ca2þ–calmodulin is able to regulate the ternary structure of actin, tropomyosin, and CaD; and subsequently affects the dynamics of actin filaments (Li et al., 2004). CaD has an elongated shape with lengths of about 74 and 53 nm for h-CaD and l-CaD, respectively. The central fragment/spacer in h-CaD (aa #208–462 of human aorta h-CaD or aa #200–446 of chick gizzard h-CaD) contains a long a-helix region with many 13–15-aa repeats. The function of this central spacer of h-CaD remains unknown but is missing in l-CaD. When microinjected into nonmuscle cells, both h-CaD and l-CaD isoforms are quickly and stably incorporated into microfilaments at stress fibers and membrane ruffles, suggesting that the central spacer is not required for actin stress fiber assembly in nonmuscle cells (Yamakita et al., 1990). The N- and C-terminal regions of CaD that are shared by both isoforms contain almost all of the known functional domains and interacting regions. The amino acid sequences of these domain regions are highly conserved throughout the evolution, as is evident in the alignment in Fig. 1.2 showing multiple alignments of amino acid sequences derived from human aorta h-CaD (M83216), human WI-38 l-CaD II (M64110), chicken brain l-CaD (M60620), chicken gizzard h-CaD (M28417), rat fibroblast l-CaD (NM_013146), and mouse fibroblast l-CaD (NM_145575).

3.1. Domains for the bindings to CaD-interacting proteins Domain mapping studies using site-directed missense and deletion mutants of recombinant CaDs as well as purified smooth muscle and nonmuscle CaDs have provided precise localizations of myosin-, actin-, tropomyosinand Ca2þ–calmodulin-binding sites as well as actomyosin ATPase inhibitory regions. Table 1.1 summarizes the positions of these functional domains on human and chicken h- and l-CaDs. Conveniently, digestion of the full-length cDNA of WI-38 l-CaD with EcoRI gives rise to two DNA fragments that encode the N-terminal fragment (namely CaD40, aa #1–243) and the C-terminal fragment (namely CaD39, aa #244–538) of human fibroblast WI-38 l-CaD II, respectively (Novy et al., 1991). CaD39 fully retains the ability of CaD to bind actin, tropomyosin, and Ca2þ–calmodulin, as well as to inhibit actomyosin ATPase activity (Eppinga et al., 2006a; Li et al., 2003b, 2004; Novy et al., 1993; Tsuruda et al., 1995). CaD39 is also capable of stabilizing actin filaments in vivo (Warren et al., 1994, 1996). The D3 fragment (aa #235–531)

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Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

MDDFERRREL MDDFERRREL ----MISRSY MDDFERRREL ----MLSRSG ----MLSGSG

RRQKREEMRL RRQKREEMRL CRQ--NLSSL RRQKREEMRL SQGRRCLATL SQGRRCLATL

EAERIAYQRN EAERIAYQRN --SKLSYQRN EAERLSYQRN --SQIAYQRN --SQIAYQRN

DDDEEEAARE DDDEEEAARE DDDEEEAARE DDDEEEAARE DDDEEEAARE DDDEEEAARE

RRRRARQERL RRRRARQERL RRRRARQERL RRRRARQERL RRRRARQERL RRRRARQERL

50 50 42 50 44 44

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

RQKQEEESLG RQKQEEESLG RQKEEGDVSG RQKEEGDVSG RQKQEEESLG RQKQEEESLG

QVTDQVEVNA QVTDQVEVNA EVTEKSEVNA EVTEKSEVNA QVTDQVEAHV QVTDQVEAHV

QNSVPDEEAK QNSVPDEEAK QNSVAEEETK QNSVAEEETK QNSAPDEESK QNSVPDEESK

TTTTNTQVEG TTTTNTQVEG RST------RST------PATANAQVEG PASSNTQVEG

DDEAAFLERL DDEAAFLERL DDEAALLERL DDEAALLERL DEEAALLERL DEEAALLERL

100 100 85 93 94 94

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

ARREERRQKR ARREERRQKR ARREERRQKR ARREERRQKR ARREERRQKR ARREERRQKR

LQEALERQKE LQEALERQKE LQEALERQKE LQEALERQKE LQEALERQKE LQEALERQKE

FDPTITDASL FDPTITDASL FDPTITDGSL FDPTITDGSL FDPTITDGSL FDPTITDGSL

SLPSRRMQND SLPSRRMQND SVPSRREVNN SVPSRREVNN SVPSRRMQNN SGPSRRMQND

TAENETTEKE TAENETTEKE VEENEITGKE VEENEITGKE SAENETAEGE SAENETAEGE

150 150 135 143 144 144

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

EKSESRQERY EKSESRQERY EKVETRQGRC EKVETRQGRC EKGESRSGRY EKRESRSGRY

EIEETETVTK EIEETETVTK EIEETETVTK EIEETETVTK EMEETEVVIT EVEETEVVIK

SYQKNDWR-D SYQKNDWR-D SYQRNNWRQD SYQRNNWRQD SYQKNSY-QD SYQKNSY-QD

AEENKKEDKE AEENKKEDKE GEEEGK--KE GEEEGK--KE AEDKKKEEKE AEDKKKEEKE

KEEEEEEKPK KEEEEEEKPK EKDSEEEKPK EKDSEEEKPK –EEEEEEKLK -EEEQE-KLK

199 199 183 191 192 191

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

RGSIGENQVE RGSIGENQ-EVPTEENQ-EVPTEENQVD GGNLGENQ-GGSLGENQ--

VMVEEKTTES ------------------VAVE-KSTDK -------------------

QEETVVMSLK ------------------EE--VVET-K -------------------

NGQISSEEPK ------------------TLAVNAE--N -------------------

QEEEREQGSD ------------------DTNAMLEGEQ -------------------

249 207 191 235 200 199

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

EISHHEKMEE ------------------SITD---AAD -------------------

EDKERAEAER ------------------KEKEEAEKER -------------------

ARLEAEERER ------------------EKLEAEEKER -------------------

IKAEQDKKIA ------------------LKAEEEKKAA -------------------

DERARIEAEE ------------------EEKQKAEEEK -------------------

299 207 191 282 200 199

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

KAAAQ----------------------KAAEERERAK -------------------

-ERERREAEE ------------------AEEEKRAAEE -------------------

RERMR-EEEK ------------------RERAKAEEER -------------------

RAAEERQRIK ------------------KAAEERERAK -------------------

-EEEKRAAEE ------------------AEEERKAAEE -------------------

341 207 191 332 200 199

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

RQRIKEEEKR ------------------RAK-AEEERK -------------------

AAEERQRIKE ------------------AAEERAK-AE -------------------

EEKRAAEERQ ------------------EERKAAEER-------------------

RARAEEEEKA -------------------AKAEKERKA -------------------

KVEEQKRNKQ -------------------AEERERAKA -------------------

391 207 191 377 200 199

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

LEEKKHAMQE ------------------EEEKRAAEEK -------------------

TKIKGEKVEQ ------------------ARLEAEKLKE -------------------

KIEGKWVNEK ------------------K---KKMEEK -------------------

KAQEDKLQTA ------------------KAQEEKAQAN -------------------

VLKKQGEEKG ------------------LLRKQEEDKE -------------------

441 207 191 424 200 199

Figure 1.2 (Continued)

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Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

TKVQAKREKL ------------------AKVEAKKESL -------------------

QED-KPTFKK ------------------PEKLQPTSKK -------------------

EEIKDEKIKK --IKDEKIKK --VKDNKVKDQVKDNKDK--IKDEKIKK --IKDEKIKK

DKEPKEEVKS DKEPKEEVKS EKAPKEEMKS EKAPKEEMKS DKEPKEEVKN DKEPKEEVKS

FMDRKKGFTE FMDRKKGFTE VWDRKRGVPE VWDRKRGVPE FLDRKKGFTE FLDRKKGFTE

490 235 218 473 228 227

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

VKSQNGE--F VKSQNGE--F QKAQNGEREL QKAQNGEREL VKAQNGE--F VKAQNGE--F

MTHKLKHTEN MTHKLKHTEN TTPKLKSTEN TTPKLKSTEN MTHKLKQTEN MTHKLKQTEN

TF--SRPGGR TF--SRPGGR AFGRS----AFGRS----AFSPSRSGGR AFSPSRSGGR

ASVDTKEAEG ASVDTKEAEG ----NLKGAA ----NLKGAA ASGD-KEAEG ASGD-KEAEG

APQVEAGKRL APQMEAGKRL NAEAGS---NAEAGS---APQVEAGKRL APQVEAGKRL

536 281 255 510 275 274

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

EELRRRRGET EELRRRRGET ---------E ---------E EELRRRRGET EELRRRRGET

ESEEFEKLKQ ESEEFEKLKQ ------KLKE ------KLKE ESEEFEKLKQ ENEEFEKLKQ

KQQEAALELE KQQEAALELE KQQEAAVELD KQQEAAVELD KQQEAALELE KQQEAALELE

ELKKKREERR ELKKKREERR ELKKRREERR ELKKRREERR ELKKKREERR ELKKKREERR

KVLEEEEQRR KVLEEEEQRR KILEEEEQKK KILEEEEQKK KVLEEEEQRR KVLEEEEQRR

586 331 290 545 325 324

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

KQEEADRKLR KQEEADRKLR KQEEAERKIR KQEEAERKIR KQEEADRKAR KQEEADRKAR

EEEEKRRLKE EEEEKRRLKE EEEEKKRMKE EEEEKKRMKE EEEEKRRLKE EEEEKRRLKE

EIERRRAEAA EIERRRAEAA EIERRRAEAA EIERRRAEAA EIERRRAEAA EIERRRAEAA

EKRQKMPEDG EKRQKMPEDG EKRQKVPEDG EKRQKVPEDG EKRQKMPEDG EKRQKMPEDG

LSDDKKPFKC LSDDKKPFKC VSEEKKPFKC VSEEKKPFKC LSEDKKPFKC LSEDKKPFKC

636 381 340 595 375 374

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

FTPKGSSLKI FTPKGSSLKI FSPKGSSLKI FSPKGSSLKI FTPKGSSLKI FTPKGSSLKI

EERAEFLNKS EERAEFLNKS EERAEFLNKS EERAEFLNKS EERAEFLNKS EERAEFLNKS

VQKSSGVKST VQKSSGVKST AQKS-GMKPA AQKS-GMKPA VQKS-GVKST VQKS-GVRST

HQAAIVSKID HQAAIVSKID HTTAVVSKID HTTAVVSKID HQAAVVSKID HQAAVVSKID

SRLEQYTSAI SRLEQYTSAI SRLEQYTSAV SRLEQYTSAV SRLEQYTNAI SRLEQYTNAI

686 431 389 644 424 423

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

EGTKSAKPTK EGTKSAKPTK VGNKAAKPAK VGNKAAKPAK EGTKASKPMK EGTKASKPMK

PAASDLPVPA PAASDLPVPA PAASDLPVPA PAASDLPVPA PAASDLPVPA PAASDLPVPA

EGVRNIKSMW EGVRNIKSMW EGVRNIKSMW EGVRNIKSMW EGVRNIKSMW EGVRNIKSMW

EKGNVFSSPT EKGNVFSSPT EKGNVFSSPG EKGNVFSSPG EKGSVFSSPS EKGSVFSAPS

AAGTPNKETA AAGTPNKETA GTGTPNKETA GTGTPNKETA ASGTPNKETA ASGTPNKETA

736 481 439 694 474 473

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

GLKVGVSSRI GLKVGVSSRI GLKVGVSSRI GLKVGVSSRI GLKVGVSSRI GLKVGVSSRI

NEWLTKTPDG NEWLTKTPDG NEWLTKTPEG NEWLTKTPEG NEWLTKSPDG NEWLTKSPDG

NKSPAPKPSD NKSPAPKPSD NKSPAPKPSD NKSPAPKPSD NKSPAPKPSD NKSPAPKPSD

LRPGDVSSKR LRPGDVSSKR LRPGDVSGKR LRPGDVSGKR LRPGDVSGKR LRPGDVSGKR

NLWEKQSVDK NLWEKQSVDK NLWEKQSVEK NLWEKQSVEK NLWEKQSVDK NLWEKQSVDK

786 531 489 744 524 523

Human Human Chick Chick Rat Mouse

h-CaD l-CaD l-CaD h-CaD l-CaD l-CaD

-VTSPTKV. -VTSPTKV. PAASSSKVTA TGKKSETNAG LRQFEKEP. PAASSSKVTA TGKKSETN-G LRQFEKEP. -VTSPTKV. -VTSPTKV.

793 538 517 771 531 530

Figure 1.2 Multiple sequence alignment of the CaD isoforms and the locations of myosin-, tropomyosin-, Ca2þ–calmodulin- and actin-binding domains. Amino acid sequences of CaD isoforms from chicken brain l-CaD (M60620), chicken gizzard h-CaD (M28417), rat fibroblast l-CaD (NM_013146), mouse fibroblast l-CaD (NM_145575), human WI-38 fibroblast l-CaD II (NM_004342–2, M64110), and

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from rat fibroblast l-CaD used in the literature (Yamashiro et al., 1995) is an equivalent form of this human CaD39. The N-terminal fragment (CaD40) of human l-CaD contains myosin binding sites (highlighted in red in Fig. 1.2) (Hemric and Chalovich, 1988; Huber et al., 1995; Ikebe and Reardon, 1988; Li et al., 2000; Marston et al., 1992; Redwood and Marston, 1993; Velaz et al., 1990; Wang et al., 1997b). The CaD39, D3, and CaD40 as well as their respective full-length l-CaD have been extensively used in the literature to investigate the in vivo function in nonmuscle cells. Tropomyosin binding regions have been detected at both N- and C-termini of CaD (Hayashi et al., 1991; Tsuruda et al., 1995; Wang et al., 1991, 1996, 1997c; Watson et al., 1990). The N-terminal tropomyosinbinding site (aa #1–152) of CaD interacts with the C-terminus (aa #142–227) of tropomyosin (Watson et al., 1990). However, this binding is readily abolished in the presence of actin (Tsuruda et al., 1995). Therefore, this site is not listed in Table 1.1. A tropomyosin binding site (highlighted in blue in Fig. 1.2) at aa #524–577 of gizzard h-CaD is predicted based on sequence similarity to the tropomyosin-binding domain of troponin T. However, this site binds skeletal muscle tropomyosin but not smooth muscle tropomyosin (Huber et al., 1995). In another experiment, a polyclonal antibody against aa #498–593 of gizzard h-CaD competed with the in vitro binding of CaD to tropomyosin, suggesting that the epitope for the antibody includes a tropomyosin-binding region (Lamb et al., 1996). Supportive of this notion, microinjection of this antibody into nonmuscle cells disrupts the stress fiber structure, further suggesting that the interaction between CaD and tropomyosin is important for microfilament bundle integrity (Lamb et al., 1996). Two other tropomyosin binding regions are defined at aa #579–635 and aa #733–742 of gizzard h-CaD (highlighted in blue in Fig. 1.2). The CaD39 fragment containing these two tropomyosin-binding sites binds tightly to both N- and C-terminal regions of the adjacent tropomyosin molecule along the actin filament, whereas the CaD40 binds very little, if at all, to the actin-tropomyosin filament, allowing this domain to project off the thin filament to interact with myosin (Tsuruda et al., 1995). This mode of interaction is further supported by the result observed from X-ray crystallography and an optical biosensor approach, which detects a strong tropomyosin binding site at aa #612–756 and two weaker tropomyosin binding sites at aa #5–50 and #180–210 of gizzard h-CaD (Hnath et al., 1996).

human aorta h-CaD (NM_033138–1) were aligned using CLUSTAL W program in Lasergene softwares (DNAstar Inc., Madison, Wisconsin). Dashes introduced represent gaps among sequences for a maximal alignment. The myosin-binding, tropomyosin-binding, and Ca2þ–calmodulin-binding are highlighted in red, blue, and yellow, respectively. The actin-binding domains and actomyosin ATPase inhibitory sites are typed in red font and underline.

Table 1.1

Locations of known functional domains in human and chicken caldesmons (CaDs)

Domains

Human aorta h-CaDa

Human WI38 l-CaD IIb

Chicken brain l-CaDc

(I) Myosin binding site

(1) 1–23 (2) 34–53 (3) 93–122 (1) 564–618 (2) 620–677 (3) 775–784 (1) 19–208 (2) 715–723

(1) 1–23 (2) 34–53 (3) 93–122 (1) 309–363 (2) 365–422 (3) 520–529 (1) 19–208 (2) 460–468

(1) 1–15 (2) 26–45 (3)78–107 (1) 269–322 (2) 324–380 (3) 478–487 (1) 13–192 (2) 418–426

(3) 747–752 (1) 707–723 within the actin-binding cluster 1, 631–722 (2) 747–756 within the actin-binding cluster 2, 749–793 (3) 775–780 within the actin-binding cluster 2, 749–793

(3) 492–497 (1) 452–468

(3) 450–455 (1) 410–426

(2) 492–501

(2) 450–450

(2) 705–714e within the actinbinding cluster 2, 707–771

(3) 520–525

(3) 478–483

(3) 733–738e within the actinbinding cluster 2, 707–771

(II) Tropomyosin binding regions (III) Ca2þ–calmodulin binding sites

(IV) Actin binding and actomyosin ATPase inhibitory sites

a b c d e

Chicken gizzard h-CaDd

(1) 1–23e (2) 34–53e (3) 86–115e (1) 524–577e (2) 579–635e (3) 733–742e (1) 19–200e (2) 673–681e site A (MWEKGNVFS) (3) 705–710e site B (NEWLTK) (1) 665–681e within the actinbinding cluster 1, 590–680.

Accession number M83216, human aorta h-CaD (793 residues) (Humphrey et al., 1992). Accession number M64110, WI-38 human fetal lung fibroblast l-CaD II (538 residues) (Hayashi et al., 1992; Novy et al., 1991). Accession number M38015 or M60620, chicken embryo brain (517 residues) (Hayashi et al., 1991); Accession number M59762, chicken gizzard l-CaD (524 residues) (Bryan and Lee, 1991). Accession number A33430, chicken gizzard h-CaD (771 residues) (Hayashi et al., 1989) and J04968 (Guo et al., 1999). Most of these domains are defined on chicken gizzard h-CaD protein using proteolytic or chemically cleaved fragments, as well as baculoviral or bacterial recombinant proteins. Based on sequence alignment (Fig. 1.2), corresponding domains in chicken l-CaD and human CaDs are deduced.

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Two strong and well-conserved Ca2þ–calmodulin binding sites (sites A and B) have been mapped to the C-terminus of CaD (highlighted in yellow in Fig. 1.2) (Bartegi et al., 1990; Fujii et al., 1987; Li et al., 2004; Lin et al., 1991; Marston et al., 1994b; Mezgueldi et al., 1994; Riseman et al., 1989; Szpacenko and Dabrowska, 1986; Wang, 2001; Wang et al., 1991; Zhan et al., 1991; Zhuang et al., 1995b). Ca2þ–calmodulin binds to CaD in an antiparallel manner (Wang et al., 1997a). Tryptophan residues within each site are crucial for the in vitro binding of CaD to Ca2þ–calmodulin (Graether et al., 1997) and for the in vivo function of human l-CaD (Eppinga et al., 2006b; Li et al., 2004). Using calmodulin affinity column chromatography with CaD fragments, another Ca2þ–calmodulin binding site has been identified to be at the N-terminus (aa #19–200) of gizzard h-CaD (Wang et al., 1989). The physiological function of this binding remains unknown. Actin binding and actomyosin ATPase inhibitory sites have been determined (red font and red underline in Fig. 1.2) (Bartegi et al., 1990; Fujii et al., 1987; Hayashi et al., 1991; Lin et al., 1991; Marston et al., 1998; Wang and Chacko, 1996; Wang et al., 1991, 1997c). The first site is located within the actin-binding cluster 1 (aa #590–680) of chicken gizzard h-CaD, determined by mass spectrometry in combination with proteolysis and actin cosedimentation, whereas the second and third sites are located within the actin-binding cluster 2 (aa #707–771) of gizzard h-CaD (Huang et al., 2003). Although an EKPA (aa #743–746) sequence from chicken gizzard h-CaD has been shown to be required for maximal inhibition of actomyosin ATPase by CaD (Wang and Chacko, 1996), the equivalent sequence is not present in the human CaD. Also, there are low affinity binding sites for actin and tropomyosin detected in the N-terminal fragment (Redwood and Marston, 1993; Tsuruda et al., 1995). The physiological significances of these sites remain unknown. Thus, they are not listed in Table 1.1. The domain responsible for inhibiting gelsolin’s severing activity has been mapped to aa #237–375 of rat l-CaD using chemically cleaved, bacterial recombinant D3 fragment (corresponding to human CaD39 fragment) (Dabrowska et al., 1996; Ishikawa et al., 1989a,b; Takiguchi and Matsumura, 2005). Based on sequence alignment, corresponding domains (aa #498–636, #244–381, #228–340, and #483–595) in human aorta h-CaD, WI-38 l-CaD II, chicken brain l-CaD, and chicken gizzard h-CaD, respectively, were deduced. This domain appears to partially overlap with the tropomyosin-binding regions on CaD. Consequently, CaD together with various tropomyosin isoforms can provide different degree of protection of actin filaments from the severing activities of gelsolin as well as different extents of annealing gelsolin-severed-actin filaments (Ishikawa et al., 1989a,b). A direct in vitro interaction between CaD and cortactin has been detected by using overlay, pull-down assay, ELISA and affinity column chromatography (Huang et al., 2006). The cortactin-binding site was

14

Jim Jung-Ching Lin et al.

further mapped to aa #693–771 of chicken h-CaD. In cultured rat aorta fibroblast cells, CaD colocalizes with cortactin to the lamellipodium and ruffles (Huang et al., 2006). Cortactin is an actin-binding protein capable of interacting with Arp2/3 complex. Through this binding at the actin branch points, cortactin stabilizes the branched actin network and regulates the dynamics of membrane ruffles, lamellipodia, and podosomes (Ammer and Weed, 2008; Weaver et al., 2002; Webb et al., 2005, 2006a, 2007; Wu and Parsons, 1993). CaD competes with cortactin for actin binding. Through this competition, CaD can attenuate Arp2/3-mediated actin polymerization (Huang et al., 2006; Yamakita et al., 2003). Although there is no evidence for direct interaction between CaD and Arp2/3 complex, CaD alone has been shown to inhibit in vitro Arp2/3-mediated actin polymerization at the nucleation step but not during the elongation process (Yamakita et al., 2003). Similar to CaD, cortactin is phosphorylated by PAK and ERK (Campbell et al., 1999; Webb et al., 2006b). The phosphorylated cortactin by PAK has a reduced affinity to actin filaments (Webb et al., 2006b). Therefore, the interaction between CaD and cortactin may provide another important step of modulating the assembly and stabilization of actin filaments in nonmuscle cells. CaD is also known to either directly or indirectly interact with fascin (an actin bundling protein) (Ishikawa et al., 1998), filamin (an actin crosslinking protein) (Nomura et al., 1987; Yonezawa et al., 1988), and cofilin (an actin depolymerizing factor) (Yonezawa et al., 1988). These interactions can, in theory, modulate their regulatory activities on the actin cytoskeleton. For example, in concert with tropomyosin isoforms, CaD effectively reduces fascin’s actin binding and bundling activities (Ishikawa et al., 1998) and filamin’s actin cross-linking activity (Nomura et al., 1987). CaD has been shown to associate with membrane phospholipids (Czurylo et al., 1993; Gusev, 2001), microtubules (tubulin) (Ishikawa et al., 1992a,b), intermediate filament components (vimentin and desmin) (Deng et al., 2007), and caltropin (the smooth muscle isoform of calcyclin, a S100 protein family member) (Gusev, 2001; Mani et al., 1992; Zhuang et al., 1995a). However, the precise domains for these interactions remain to be determined. The association of CaD with many of these proteins and phospholipids is affected by the presence of Ca2þ–calmodulin and by way of phosphorylation by various kinases. In other words, CaD activity can be effectively regulated in cells by multiple mechanisms.

3.2. Phosphorylated residues on CaD by various kinases Both h-CaD and l-CaD were found to be phosphorylated during smooth muscle contraction and migration upon stimulation, during nonmuscle cell division, spreading and migration, during cell transformation by oncogenic

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v-erbB, or during cell infection with Kaposi sarcoma-associated herpes virus (Adam and Hathaway, 1993; Adam et al., 1992; Boerner et al., 2003; Bogatcheva et al., 2006; Cuomo et al., 2005; Gerthoffer et al., 1996; Hai and Gu, 2006; Kordowska et al., 2006b; Matsumura and Yamashiro, 1993; Morgan and Gangopadhyay, 2001). The kinases identified so far include Cdc2 kinase (B-cyclin/Cdk1), B2-cyclin/Cdk1, K-cyclin/Cdk6, PAK, ERK, protein kinase C (PKC), Ca2þ–calmodulin kinase II (CaMKII), casein kinase II (CKII), and v-erbB tyrosine kinase. Table 1.2 summarizes known phosphorylation sites on human and chicken CaD isoforms by these kinases and the resulting effects on CaD’s functions/properties. Relevant references for each kinase are also listed in Table 1.2. In general, phosphorylation of CaD by ser/thr kinases reduces its binding properties to actin filament, tropomyosin, Ca2þ–calmodulin, or myosin and/or releases the inhibition of actomyosin ATPase activity, depending on the location of phosphorylation sites. In contrast, tyrosine phosphorylation of CaD by oncogenic v-erbB increases its binding to signal adaptor protein, Grb2, and promotes the formation of a transformation-associated signaling complex, leading to stress fiber disassembly, anchorage-independent cell proliferation, and transformation.

3.3. CaD and calponin interaction It is well-established that smooth muscle and nonmuscle cell contraction is regulated by the Ca2þ–calmodulin-dependent phosphorylation of the myosin regulatory light chain (MRLC) by myosin light chain kinase (MLCK) (so-called the myosin-linked regulation) (Adelstein and Eisenberg, 1980; Kohama et al., 1996; Moussavi et al., 1993). However, accumulating evidence suggest that the interaction of myosin with actin in these cells can be also regulated by the actin- and Ca2þ–calmodulin-binding proteins, CaD (Sobue and Sellers, 1991), and calponin (Takahashi and Yamamura, 2003) (so-called the actin-linked regulation). As summarized in Table 1.3, both CaD and calponin have very similar properties in inhibiting the actinactivated ATPase activity and the in vitro actin filament movement on myosin. Although the modes of inhibition by these proteins are clearly different, this inhibition can generally be reversed by the addition of Ca2þ– calmodulin and through phosphorylation by various kinases (Table 1.3). Both CaD and calponin are also capable of interacting with many actinbinding proteins, phospholipids, microtubules, and intermediate filaments, suggesting that they may play a role in remodeling the cytoskeleton for shape changes, in addition to modulating actin-myosin interaction for force development and cell migration. The intracellular localizations of these two proteins are partially overlapped and complementary to each other at both contractile and cytoskeletal domains, further suggesting that they may

16

Table 1.2 Known phosphorylation sites and effects on CaD activity Kinases

(I) Cdc2 kinase (B-cyclin/ Cdk1)

(II) p21-activated kinase (PAK)

(III) Extracellular signalregulated kinase (ERK) and p38 MAPK

(IV) Protein kinase C (PKC) (V) Ca2þ-calmodulin kinase II (CaMKII)

Phosphorylation sites

Effects 638

724

730

753

759

Human aorta h-CaD: T , S , T , T , S , S789 Human WI-38 l-CaD II: T383, S469, T475, T498, S504, S534 Chicken brain l-CaD: T230 (avian only), S342, S427, T433, T456, S462 Chicken gizzard h-CaD: T485 (avian only), S597, S682, T688, T711, S717 Human aorta h-CaD: S714, S744 Human WI-38 l-CaD II: S459, S489 Chicken brain l-CaD: S417, S447 Chicken gizzard h-CaD: S672, S702 Human aorta h-CaD: S759, S789 Human WI-38 l-CaD II: S504, S534 Chicken brain l-CaD: S462 Chicken gizzard h-CaD: S717 Human aorta h-CaD: S643, S456, S783 Human WI-38 l-CaD II: S388, S401, S528 Chicken brain l-CaD: S347, S360, S486 Human aorta h-CaD: S73, S643, S677, S783 Human WI-38 l-CaD II: S73, S388, S422, S528

(1) # binding to actin (2) no effect on binding to calmodulin (3) release inhibition of actomyosin ATPase activity (1) # binding to actin (2) # binding to calmodulin (3) release inhibition of actomyosin ATPase activity (1) " isometric tension in smooth muscle (2) # binding to actin, " actin stress fiber disassembly (3) release inhibition of actomyosin ATPase activity (1) # binding to actin (2) # binding to calmodulin (3) partially release inhibition of actomyosin ATPase activity (1) # binding to myosin (2) # binding to actin (3) release inhibition of actomyosin ATPase activity

Chicken brain l-CaD: S18 (avian only), S51 (avian only), S65, T229(avian only), S235 (avian only), S347, S380, S486 Chicken gizzard h-CaD: S26 (avian only), S59 (avian only), S73, T484(avian only), S490 (avian only), S602, S635, S741 (VI) Casein kinase II (CKII) Human aorta h-CaD; S73, T83

(VII) v-erbB tyrosine kinase

Human WI-38 l-CaD II: S73, T83 Chicken brain l-CaD: S18 (avian only), S65, T75 Chicken gizzard h-CaD: S26 (avian only), S73, T83 Human aorta h-CaD: Y27, Y682 Human WI-38 l-CaD II: Y27, Y427 Chicken brain l-CaD: Y19, Y385 Chicken gizzard h-CaD: Y27, Y640

(1) # binding to myosin and preventing tethering of myosin to actin filaments (control tonic contraction) (2) # binding to tropomyosin

(1) " binding to Shc–Grb2–Sos complex (" transformation-associated signaling complex) (2) stress fiber loss

17

(I) Cdc2 kinase: (Li et al., 2003b; Mak et al., 1991; Yamashiro et al., 1990, 1991, 2001); Mitosis-specific phosphorylation (proline-directed) sites determined in rat l-CaD are S249 (rodent only), S462, T468, S491, S497, and S527. There is another putative site at S377 by Cdc2 kinase but not detected in this assay (Matsumura and Yamashiro, 1993). The equivalent sites listed in this Table for human and chicken CaDs are deduced from these residues except S249 of rat l-CaD. (II) PAK: (Eppinga et al., 2006ab; Foster et al., 2000; McFawn et al., 2003; Van Eyk et al., 1998); PAK induces Ca2þ-independent contraction of skinned muscle fibers from guinea pig taenia coli and enhances phosphorylation of h-CaD but not myosin regulatory light chain (MRLC) (Van Eyk et al., 1998). PAK-mediated changes in the organization of actin cytoskeleton regulate cell morphology and motility in most eukaryotic cells (Sells, 1999; Sells et al., 1999) and breast cancer cells (Adam et al., 2000). Reversible CaD phosphorylation at PAK-responsive sites is required for normal cell migration (Eppinga et al., 2006a) and cytokinesis (Eppinga et al., 2006b). (III) ERK and p38 MAPK: (Adam and Hathaway, 1993; Childs et al., 1992; D’Angelo et al., 1999; Hedges et al., 1998, 2000; Huang et al., 2003; Kordowska et al., 2006a); Both Cdc2 kinase and MAPK (ERK and p38) recognize similar proline-directed serine/threoine residues on CaD but have different preferred sites (Hedges et al., 1998). These ERK and p38 MAPK phosphorylation sites correspond to S497 and S527 in the rodent fibroblast l-CaD.

18 Table 1.2 (Footnote continued ) (IV) PKC: (Adam et al., 1992; Ikebe and Hornick, 1991; Matsumura and Yamashiro, 1993; Tanaka et al., 1990; Umekawa and Hidaka, 1985; Vorotnikov et al., 1994). These PKC phosphorylation sites are derived from equivalent sites defined in vitro on porcine stomach h-CaD (Adam et al., 1992; Ikebe and Hornick, 1991). However, in phorbol ester (direct and potent PKC activator)-treated intact smooth muscle, h-CaD is phosphorylated at sites different from those in vitro PKC phosphorylation sites. The in vivo sites are now known to be ERK phosphorylation sites (Adam et al., 1992). In rabbit colonic smooth muscle, acetylcholine induces a significant association of hCaD with PKCa and sustained phosphorylation of h-CaD at Ser789 (one of ERK phosphorylation sites). Acetylcholine also induces phosphorylation of HSP27 and enhances the association of phospho-CaD and phospho-HSP27. As a result, phospho-CaD dissociates from tropomyosin and leading to the interaction between actin and myosin (Somara and Bitar, 2006). (V) Ca2þ-calmodulin kinase II (CaMKII): (Hemric et al., 1993; Ikebe and Reardon, 1990; Ngai and Walsh, 1987; Sutherland and Walsh, 1989); Smooth muscle CaMKII tightly associates with h-CaD. The preferred phosphorylation sites are located in the N-terminal myosin binding domain, whereas slower phosphorylation occurs in the C-terminus containing Ca2þ-calmodulin and actin-binding domains (Ikebe and Reardon, 1990). In vivo evidence to support the involvement of CaMKII phosphorylation of CaD is still scarce. In bovine pulmonary artery endothelial cells, thrombin induced endothelial cell barrier dysfunction, which had been shown to involve the activation of both CaMKII and ERK. However, the phosphorylation of l-CaD in vivo has been found to be at least at S534 (equivalent to S789 on h-CaD), which is different from those in vitro CaMKII sites (Borbiev et al., 2003). (VI) Casein kinase II (CKII): (Bogatcheva et al., 1993; Sutherland et al., 1994; Wang and Yang, 2000; Wawrzynow et al., 1991); Phosphorylation sites by CKII are located outside of the myosin-binding sites, suggesting the decrease in the myosin binding affinity may be a result of conformational changes after CKII phosphorylation (Wang and Yang, 2000). (VII) v-erbB tyrosine kinase: (Boerner et al., 2000, 2003a; McManus et al., 1997, 2000; Wang et al., 1999); The tyrosine phosphorylation sites are initially determined on chicken gizzard l-CaD at Y27 and Y393 (Wang et al., 1999), which are adjacent to major myosin and actin binding domains. Formation of transformation-associated signaling complex in v-erbB (EGF receptor mutant) transformed cells requires the phosphorylation at the tyrosine residues of CaD (McManus et al., 1997; Wang et al., 1999). This signaling complex composed of the signal adaptor proteins (Shc, Grb2, Nck, and Sos), and tyrosine phosphorylated forms of PAK, CaD, and myosin light chain kinase (MLCK) provides a mechanism regulating actin stress fiber disassembly in transformed cells (Boerner et al., 2003b; McManus et al., 2000). After tyrosine phosphorylations of these substrates, CaD can then mediate the interaction with SH2- and SH3-containing protein Grb2, and PAK can be constitutively activated, whereas MLCK decreases its catalytic activity (Boerner et al., 2003b). The functional consequence of these modifications results in cytoskeletal alterations (such as stress fiber loss and altered adhesion) and anchorage-independent cell growth and transformation (Boerner et al., 2003a). The ligand-independent oncogenic signaling by v-erbB is different from the ligand-dependent mitogenic signaling by EGF receptor in normal cells.

Table 1.3 Property comparisons between CaD and calponin Property

CaD

Calponin

(I) Protein isoforms and tissue expression pattern

Single gene encodes h- CaD (in differentiated smooth muscle) and l-CaD (in all cell types except skeletal muscle) via alternative splicing

(II) Bindings to myosin, actin, tropomyosin, and Ca2þ–calmodulin

Yes for both h- and l-CaD

Three distinct genes encode h1-calponin (in differentiated smooth muscle, developing heart, skeleton, and undifferentiated osteoblasts), h2-calponin (in both smooth muscle and nonmuscle cells, such as human epidermal keratinocytes, fibroblasts, lung alveolar cells, endothelial cells, and white blood cells) and acidic h3-calponin (in smooth muscle, lung, kidney, stomach, and brain) ( Jin et al. (2008); Wu and Jin (2008)). Yes for h1-calponin (Childs et al. (1992); Szymanski and Goyal (1999); Szymanski and Tao (1993); Takahashi et al. (1988)) and likely for h2- and acidic calponin (Wu and Jin (2008)). Yes (Fujii et al. (2000); Mabuchi et al. (1997); Wang and Gusev (1996)) Yes (Fujii et al. (1997); Fujii and Koizumi (1999)) Yes (Bogatcheva and Gusev (1995a,b); Gusev (2001)) Yes for h1-calponin ( Jin et al. (2008); Rozenblum and Gimona (2008)) and likely for h2- and acidic calponin (Wu and Jin (2008)). No effect (Winder and Walsh (1990))

19

(III) Binding to intermediate Yes (Deng et al. (2007)) filaments (IV) Binding to microtubules Yes (Ishikawa et al. (1992a,b)) (V) Binding to phospholipids Yes (Bogatcheva et al. (1994); Gusev (2001)) Yes for both h- and l-CaD (Marston (VIa) Inhibition of and Redwood (1991); Sobue and actomyosin ATPase Sellers (1991)) activity (VIb) Tropomyosin effect on Tropomyosin is necessary for full this inhibition inhibition (Chalovich et al. (1987); Smith et al. (1987))

(continued)

20

Table 1.3 (continued) Property

CaD

Calponin

(VIc) Reversal of the inhibition by Ca2þ–calmodulin (VId) Reversal by phosphorylation (VIIa) Inhibition of in vitro actin filament movement

Yes for both h-and l-CaD (Marston and Redwood (1991); Sobue and Sellers (1991)) Yes for both h- and l-CaD in vitro and in vivo Yes, graded inhibition (for h-CaD). Tropomyosin reduces the amount needed for inhibition. (Haeberle et al. (1992); Shirinsky et al. (1992)) modulation of the apparent on-rate for cross bridges (Shirinsky et al. (1992)) Yes (Shirinsky et al. (1992))

Yes (Abe et al. (1990))

(VIIb) mode of inhibition (VIIc) Reversal of the inhibition by Ca2þ–calmodulin (VIII) Phosphorylation (a) In Both h- and l-CaD are phosphorylated vitro by various kinases (see Table 1.2)

(b) In vivo during smooth muscle contraction

Yes (see Table 1.2)

Yes (Winder and Walsh (1990)) Yes, ‘‘all or none’’ type of inhibition (for h1-calponin). Tropomyosin has no effect on the amount of calponin needed for inhibition. (Haeberle and Hemric (1994); Shirinsky et al. (1992))modulation of the apparent off-rate for cross bridges (Shirinsky et al. (1992)) Yes (Shirinsky et al. (1992))

The h1-calponin is phosphorylated by PKC (Nakamura et al. (1999)), CaMKII (Winder et al. (1993)) and Rho-kinase (Kaneko et al. (2000)). The effects of phosphorylation lead to a weaker binding to actin and a reversal of inhibiting actomyosin ATPase. Not detectable (Adam et al. (1995); Barany and Barany (1993); Barany et al. (1991); Gimona et al. (1992)). Calponin directly binds to ERK and PKC but is not phosphorylated, cotranslocates with

(IX) Localization (a) in smooth muscle cells

The h-CaD preferentially localizes to thin filaments that are near myosin filaments (contractile or actomyosin domain) but the distribution of CaD is narrower than tropomyosin (Furst et al. (1986); Mabuchi et al. (1996, 2001))

these kinases to the plasma membrane and activates PKC upon agonist-stimulation of smooth muscle contraction, suggesting a role for calponin as a signaling molecule (Morgan and Gangopadhyay (2001)). The h1-calponin localizes to both contractile and cytoskeleton domains but more concentrates in the cytoskeleton, additionally localizes in the cytoplasmic dense bodies and in the adhesion plaques near the cell surface.

Involves in bridging various cytoskeletal proteins and/or interfacing the cytoskeleton and contractile apparatus. (Mabuchi et al. (1996); North et al. (1994)) The colocalization of h2-calponin with nonmuscle tropomyosins to actin stress fibers and cytoskeleton. The specific localization of h2-calponin around the nuclei of dividing cells (Hossain et al. (2006)) The h3-calponin mostly restricts to neuronal tissues and accumulates in glia cells and astrocytes of the brain cortex, hippocampus and cerebellum, and in the choroid plexus (Plantier et al. (1999); Represa et al. (1995)). 21

(continued)

Table 1.3 (continued) Property

CaD

Calponin

(IX) Localization (b) in nonmuscle cells

The l-CaD localizes to stress fibers, membrane ruffles, and lamellipodia (Bretscher (1986); Dabrowska et al. (2004); Huber (1997); Matsumura and Yamashiro (1993); Sobue and Sellers (1991); Sobue et al. (1988))

(X) Targeted deletion in mice

h-CaD-null mice survive with up-regulation of l-CaD in phasic smooth muscles, such as bladder and intestine. There are no reported changes in smooth muscle contractile properties from h-CaD-null mice (Guo and Wang (2005)).

Cnn1-null mice show increase in unloaded shortening velocity of smooth muscle contraction, display early onset of cartilage formation and ossification, accelerate healing of bone fractures, and impair vessel maturation (Matthew et al. (2000); Taniguchi et al. (2001); Yamamura et al. (2007); Yoshikawa et al. (1998)). The h2-calponin knockdown zebrafish embryos exhibit many circulation defects, enlarged cerebral ventricles and pericardial edema (Tang et al. (2006)).

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cooperatively and synergistically modulate actin–myosin interaction and cytoskeleton dynamics. In support of this notion, the direct interaction between calponin and CaD has been observed from two different laboratories (Graceffa et al., 1996; Vancompernolle et al., 1990), although another laboratory could not detect this interaction (Czurylo et al., 1997). The physiological significance of the CaD-calponin interaction remains unknown. Both CaD and calponin were shown to be involved in histamine-induced sustained contraction of arterial smooth muscle in vivo (Barany et al., 1992). In vitro, CaD and calponin bind simultaneously to actin filaments when used in sub-saturating amounts, whereas CaD and calponin directly compete for binding to actin when used in excess amounts (Makuch et al., 1991). These results suggest that they may work together in regulating actomyosin ATPase activity and actin cytoskeletal dynamics. It is also known that the mode of inhibition of actomyosin ATPase activity and the requirement of tropomyosin for this inhibition are different between CaD and calponin (Table 1.3). In vitro, CaD has no significant influence on the calponin-induced inhibition of actomyosin ATPase activity and vise versa (Winder et al., 1992). The h1-calponin (Cnn1) knockout mice show neither overexpression nor ectopic expression of h2 and h3-calponin (Yoshikawa et al., 1998) but exhibit altered contractile properties in smooth muscles (Matthew et al., 2000). Interestingly, the h-CaD from the Cnn1 knockout mouse smooth muscles shows a significantly decreased electrophoretic mobility, which may reflect a change in phosphorylation or a different CaD variant (Yoshikawa et al., 1998). It remains to be determined whether this variant of h-CaD may compensate for the loss of h1-calponin in terms of modulating smooth muscle contraction (Matthew et al., 2000). On the other hand, the h-CaD specific knockout mice exhibit no obvious phenotype in smooth muscle tissues but have a significant increase in l-CaD in phasic smooth muscles (Guo and Wang, 2005). The expression of calponin isoforms in the h-CaD knockout mice remains to be determined. Several lines of evidence from recent studies have demonstrated a novel regulation of calponin expression and degradation by mechanical tension in the actin cytoskeleton (Hossain et al., 2005, 2006). In epidermal keratinocytes, lung alveolar cells, and fibroblasts cultured on polyacrylamide gels of different stiffness, the expression of h2-calponin is affected by the stiffness of the culture substrate. When cells are cultured on soft gel that applies less traction force to the cell and, therefore, lower mechanical tension in the cytoskeleton, the level of h2-calponin is significantly lower than that in cells cultured on a hard gel or a rigid plastic dish. Lowering the tension of actin cytoskeleton by inhibiting nonmuscle myosin ATPase decreased h2-calponin expression. Prolonged deflation of lung led to a rapid degradation of h2calponin, which was prevented by inflation of the lung to the in situ expanded

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volume. Decreasing mechanical tension in cultured alveolar cells by reducing the dimension of culture matrix reproduced the degradation of h2-calponin. Inhibition of myosin ATPase also resulted in the degradation of h2calponin in alveolar cells, showing a determining role of the tension in the actin cytoskeleton. The tension regulation of h2-calponin synthesis and degradation demonstrates a novel mechanical regulation of cellular biochemistry. Considering the established interrelationship between CaD and calponin, it will be interesting to investigate whether the gene expression and posttranslational regulation of CaD are also modulated by mechanical tension in the actin cytoskeleton.

4. CaD and Regulation of Smooth Muscle Contraction In vitro characterization implies that h-CaD plays important roles in modulating smooth muscle contraction. In support of this notion, h-CaD is found to localize within the subset of actin filaments that are also associated with myosin (the actomyosin domain) (Furst et al., 1986). Incubation of permeabilized smooth muscle cells with exogenous CaD prolongs the rate of muscle relaxation (Albrecht et al., 1997), while depletion of h-CaD by antisense oligodeoxynucleotides results in the generation of contractile force in resting vascular smooth muscle (Earley et al., 1998). The proposed involvement of h-CaD in mediating the regulation of the actin-myosin interaction and the inhibition of actomyosin ATPase activity is further supported by several lines of in vivo evidence. A synthetic peptide (GS17C, aa #666–682 of chicken gizzard h-CaD) overlapping with the actin- and Ca2þ–calmodulin-binding sites (Zhan et al., 1991) has been shown to induce a sustained elevation of basal contractile tone in permeabilized, ferret aorta and portal vein cells (Katsuyama et al., 1992). It is likely that this peptide competes with endogenous CaD, displays part of the CaD molecule away from actin filaments, and reverses the CaD’s inhibitory action. Indeed, photo-crosslinking and fluorescence quenching experiments show that GS17C peptide dissociates the C-terminal region of CaD from actin in a manner similar to ERK-mediated phosphorylation (Huang and Wang, 2006). Similarly, two synthetic peptides (IK29C, aa #1–27 and MY27C, aa #25–53) spanning the myosin-binding domain of CaD are able to induce a dose-dependent basal tone of permeabilized smooth muscle cells and to specifically prevent agonist-induced contractions. These results imply that the N-terminal myosin-binding domain of CaD that tethers actin

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to myosin is essential for CaD’s inhibitory function. Disruption of this tethering by either peptide treatment induces basal tone and prevents agonist-induced contraction (Lee et al., 2000). The addition of Ca2þ–calmodulin reverses the in vitro CaD’s inhibitory actions. There is not very much in vivo evidence except for the GS17C antagonist peptide experiment to directly support Ca2þ–calmodulin regulating CaD activity in smooth muscle cells. However, considering the high binding affinity of calmodulin to CaD (Zhuang et al., 1996) and the local calmodulin concentration increase in response to agonist stimulation, the CaD activity regulated by the binding of calmodulin may actually take place in living smooth muscle cells (Hulvershorn et al., 2001). Controversial results have been obtained from many in vivo experiments performed to address whether the CaD activity in smooth muscle cells is regulated by phosphorylation. Many kinases can phosphorylate CaD in vitro and their phosphorylation residues have been determined (Table 1.2). The phorbol esters are well-known activators of PKC to stimulate many types of smooth muscles. The phosphorylation sites in CaD determined from phorbol ester-stimulated intact canine aorta are different from the in vitro phosphorylation sites of h-CaD by PKC (Adam et al., 1992). Instead, the in vivo sites (S759 and S789 of human aorta h-CaD) identified from phorbol ester-treated smooth muscle are the same sites for ERK MAPK, p38MAPK, and Cdc2 kinase (Adam and Hathaway, 1993; Childs et al., 1992; Hedges et al., 1998; Yamboliev et al., 2000). Phosphorylation site-specific antibodies detect phosphorylation only at S789 but not S759 of h-CaD in vascular, airway, and colonic smooth muscles (Cook et al., 2000; D’Angelo et al., 1999; Hedges et al., 2000; Yamboliev et al., 2000). Moreover, ERK activity and CaD phosphorylation follow a time course similar to that of various agonist-induced contractions (Dessy et al., 1998; Franklin et al., 1997; Gerthoffer et al., 1997), supporting the role of CaD phosphorylation in regulating smooth muscle contraction. It is known that muscarinic M2 receptor signaling activates both ERK and p38 MAPK (Gerthoffer, 2005), which are known to phosphorylate the same sites on h-CaD in vitro (Hedges et al., 1998), and leads to smooth muscle contraction, cell migration, and cytoskeletal remodeling (Gerthoffer, 2005). With the use of kinase specific inhibitors, it is possible to distinguish the roles of these kinases on CaD phosphorylation and muscle contraction (Hedges et al., 2000; Yamboliev et al., 2000). From these inhibitor studies on canine pulmonary artery smooth muscle fibers, it is clear that ERK MAPK but not p38 MAPK is the major CaD kinase in vivo. However, neither the activation of ERK nor phosphorylation of h-CaD are necessary for force development, which instead is modulated by the p38 MAPK and HSP27 phosphorylation pathway (Yamboliev et al., 2000). In cultured

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pulmonary artery smooth muscle cells expressing a high amount of l-CaD with minor h-CaD, the activation of ERK and l-CaD phosphorylation is, instead, required for modulating the platelet derived growth factor (PDGF)stimulated cell migration (Yamboliev and Gerthoffer, 2001). On the other hand, the activation of p38MAPK and the phosphorylation of nonmuscle l-CaD are essential for migration of tracheal smooth muscle cells in response to urokinase type plasminogen activator (agonist) (Goncharova et al., 2002). Therefore, the interpretation and comparison of experimental results reported in the literature must carefully consider agonist-, culture-, animal species-, and muscle tissues-specific effects. Nevertheless, recent studies with freshly isolated colonic smooth muscle cells stimulated with acetylcholine reveal a novel role for CaD phosphorylation at S789 in regulating actomyosin interaction and contraction (Somara and Bitar, 2006). Acetylcholine induces PKC-mediated smooth muscle contraction through the activation of ERK and CaD phosphorylation at S789 as well as the activation of p38 MAPK and HSP27 phosphorylation (Gerthoffer, 2005). Somara and Bitar further demonstrated that acetylcholine treatment of colonic smooth muscle cells induces an increase in phosphorylation of HSP27, which together with PKCa translocates to the particulate/membrane fraction where PKCa is phosphorylated. Phospho-PKCa, on translocation, binds to nonphosphorylated CaD and tropomyosin, resulting in phosphorylation of CaD by activated ERK and p38 MAPK. The phospho-CaD dissociated from phospho-PKCa can then bind to phospho-HSP27 resulting in a conformational change of CaD, partial dissociation from actintropomyosin, and releasing the CaD’s inhibitory actions on actomyosin interaction and muscle contraction (Somara and Bitar, 2006). PAK is another kinase able to induce Ca2þ-independent contraction in skinned smooth muscle fibers from guinea pig tenia coli and from dog trachealis with concomitant increase in phosphorylation of h-CaD (McFawn et al., 2003; Van Eyk et al., 1998). Although PAK can also phosphorylate MRLC in vitro, the level of MRLC phosphorylation in these contracted muscles remains similar to that in relaxed muscle fibers. Subsequent biochemical studies identified two phosphorylation sites for PAK on the C-terminus of CaD, which are different from those sites phosphorylated by other known kinases (Table 1.2). Unlike phosphorylation of CaD by Cdc2 kinase, PAK phosphorylation of CaD leads to only a moderate reduction in its binding affinity for actin filaments. However, it significantly reduces the ability of CaD to inhibit actomyosin ATPase activity (Foster et al., 2000). PAK may also have a significant role in cultured dog tracheal smooth muscle cell migration and airway remodeling by signaling to p38 MAPK and other pathways (Dechert et al., 2001).

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However, the relevance of PAK phosphorylation of CaD to smooth muscle cell migration remains to be determined.

5. CaD and Regulation of Nonmuscle Cell Motility and Cytoskeleton Dynamics In contrast to the highly ordered structure of actin thin filaments in muscle cells, the actin structures in nonmuscle cells are extremely dynamic. In this context, regulation of actin cytoskeleton dynamics has become a profound function for the nonmuscle l-CaD isoform, in addition to regulation of actomyosin ATPase activity. Biochemical studies have demonstrated that l-CaD inhibits the actin-severing activity of gelsolin and enhances the reannealing of gelsolin-severed actin filaments (Dabrowska et al., 1996; Ishikawa et al., 1989a,b; Takiguchi and Matsumura, 2005). On the other hand, l-CaD or its C-terminal fragment CaD39 can potentiate the binding of nonmuscle tropomyosin to actin filaments (Novy et al., 1993; Yamashiro-Matsumura and Matsumura, 1988). Nonmuscle l-CaD competes with fascin for actin binding sites and thus inhibits actin bundling activity of fascin in a Ca2þ–calmodulin-sensitive manner (Ishikawa et al., 1998). Therefore, it seems that, while CaD stabilizes the filamentous structures, it also prevents further assembly of actin filaments into bundles. Furthermore, l-CaD can inhibit Arp2/3 complex-mediated actin polymerization in vitro (Yamakita et al., 2003). This Arp2/3 complex-mediated actin branching process plays an important role in regulating the dynamics of motile structures such as membrane ruffles and lamellipodia. Although it has not been shown with l-CaD, gizzard h-CaD stimulates actin polymerization and induces actin polymerization from profilin-G-actin complex, which is also enriched in membrane ruffles and lamellipodia (Galazkiewicz et al., 1989, 1991). Purified l-CaD or its recombinant fragment inhibits actomyosin ATPase activity and the inhibition is reversed by the addition of Ca2þ– calmodulin and/or by phosphorylation (Eppinga et al., 2006a; Huang et al., 2003; Li et al., 2004; Sobue et al., 1988; Yamashiro et al., 1995). Because both h-CaD and l-CaD are derived from the same gene of the same species and because the structure and sequence are highly conserved among all CaDs from various species (Czurylo, 2000), most evidence obtained from the in vitro studies with h-CaD are likely applied to l-CaD. For example, CaD is able to decrease the binding of filamin to actin filaments, preventing actin filaments from cross-linking (Nomura et al., 1987). CaD can prevent actin filaments from depolymerization and the binding of actin depolymerizing factor cofilin can dissociate CaD from actin filaments (Yonezawa et al., 1988).

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Together with the recent finding that CaD interacts with cortactin (Huang et al., 2006), the results obtained from these in vitro studies suggest a functional significance for l-CaD in regulating actin cytoskeleton dynamics.

5.1. Intracellular localization of l-CaD and its functions In interphase cells, l-CaD is localized to stress fibers and at the leading lamellipodia and membrane ruffles, as well as at the podosomes of certain highly motile normal cells and cancer cells (Bretscher and Lynch, 1985; Lin et al., 1988; Nakamura et al., 1993; Owada et al., 1984; Tanaka et al., 1993; Yamakita et al., 1990). The stress fibers are made of actin microfilament bundles containing myosin, tropomyosin, a-actinin, and others required for force production. Two functionally and spatially distinguished types, central (ventral and dorsal) and peripheral, of stress fibers are observed in fibroblasts (Small et al., 1998). The lamellipodium and membrane ruffle are the dynamic structures of migrating cells and also contain myosin and low molecular mass tropomyosin (Fukui et al., 1989; Gunning et al., 2007; Kolega, 1998; Lin et al., 1988, 1997, 2008). Such colocalization of CaD with the contractile proteins is in agreement with the concept that CaD is a regulatory protein for the actomyosin system. In normal cells, CaD is absent in focal adhesion, at where the stress fibers terminate (Tanaka et al., 1993). Upon transformation by Rous sarcoma virus or by tumor promoters, cells lose their stress fibers and rearrange their relatively immobile focal adhesions into highly dynamic podosomes (Nakamura et al., 1993; Tanaka et al., 1993), which contain short actin bundles as core domain, perpendicular to the substratum and surrounded by microfilaments (ring domain) containing adhesion mediators such as vinculin, talin, paxillin, and kinases (Buccione et al., 2004; Linder and Aepfelbacher, 2003; Linder and Kopp, 2005). CaD together with actin regulators such as cortactin, Arp2/3 complex, WASP, and Cdc42 is found in the core domain of podosomes, whereas myosin, a-actinin, and low molecular mass tropomyosin are associated with the ring structure (Hilenski et al., 1991; Stickel and Wang, 1987; Tanaka et al., 1993). Podosomes are originally found in macrophages, osteoclasts, eosinophils, cultured cardiomyocytes, and transformed fibroblasts (Linder and Kopp, 2005; Osiak et al., 2005). Recently, it is also shown that endothelial cells and vascular smooth muscle cells can be induced to form podosomes in responses to transforming growth factor-b (Varon et al., 2006), phorbol esters (Burgstaller and Gimona, 2005; Eves et al., 2006; Hai et al., 2002), or Cdc2 activation (Moreau et al., 2003, 2006; Osiak et al., 2005). Podosomes are thought to function as sites of cell adhesion and active extracellular matrix remodeling for cell migration and invasion, because the core domain contains metalloproteases (MT1-MMP and MMP-9) and acts as sites of MMP secretion (Chen, 1989; Chen et al., 1984, 1985; Ochoa et al., 2000). Macrophages from patients with Wiskott–Aldrich syndrome

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(WAS) cannot properly form podosomes due to defective in WAS protein (WASP) and these WAS macrophages are also found to be defective in cell motility (Linder et al., 1999). The l-CaD found in the core of podosomes of phorbol ester-stimulated smooth muscle cells has been shown to play a significant role in regulating podosome formation (Eves et al., 2006). The ability of l-CaD redistribution in responses to physiological changes within cells also implies the involvement of l-CaD in controlling actin filament reorganization for specific functions. These lines of in vivo evidence conclude that CaD functions in ligand-induced receptor capping of lymphocytes (Mizushima et al., 1987; Walker et al., 1989), in catecholamine secretion of stimulated adrenal chromaffin cells (exocytosis) (Burgoyne et al., 1986), in podosome formation of many transformed cells (Nakamura et al., 1993; Tanaka et al., 1993), in stabilizing stress fibers for the control of hormone release from pituitary cells (Castellino et al., 1992, 1995; Janovick et al., 1991), and in contractile ring of mitotic cells (Eppinga et al., 2006b; Li et al., 2003b; Yamashiro et al., 1990, 1991). Microinjection of anti-CaD monoclonal antibody C21, which has been shown to inhibit the binding of CaD to Ca2þ–calmodulin and to actin filaments (Lin et al., 1991), results in a drastic reduction of instantaneous speed and saltation distance of intracellular granule movement (Hegmann et al., 1991). A similar effect on granule movement has been observed in experiments with anti-tropomyosin antibody injection (Hegmann et al., 1989). These results suggest that l-CaD and tropomyosin participate in the regulation of actomyosin system involved in the intracellular granule movement. Similarly, microinjection of anti-CaD polyclonal antibody into nonmuscle cells disrupts microfilament bundles and this antibody is shown to be able to compete with tropomyosin to bind to CaD (Lamb et al., 1996). Although the effect of antibody on intracellular granule movement has not been addressed in this experiment, these results further suggest the importance of CaD-tropomyosin interaction in actin filament integrity and function in vivo. During platelet activation, CaD through its myosin-binding domain binds to soluble myosin and then facilitates the translocation of myosin to the insoluble actin cytoskeleton (Hemric et al., 1994). This translocation together with MRLC phosphorylation allows platelet undergoing shape change, cell aggregation, granule secretion, and clot retraction. Upon phorbol ester-treatment of endothelial cells, the rearrangement of actin cytoskeleton concomitant with the loss of endothelial barrier function (shape change) is shown to be associated with increased l-CaD phosphorylation but not with MRLC phosphorylation (Bogatcheva et al., 2006; Moy et al., 2004). Although the phosphorylation sites have not been completely determined, at least the S534 site on human l-CaD by ERK1/2 is involved (Bogatcheva et al., 2006). Similarly, thrombin-induced endothelial cell barrier dysfunction (shape change) also involves a combination of CaD

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phosphorylation at S534, actin cytoskeletal rearrangement, and contraction. At least three kinase (CaMKII, ERK, and p38 MAPK) activities are needed to modulate this MLCK-independent regulation of thrombin-induced endothelial cell barrier dysfunction (Borbiev et al., 2003, 2004). Therefore, CaD plays important role in regulating endothelial cell barrier function through the actin cytoskeletal remodeling.

5.2. Regulation of l-CaD activity in nonmuscle cells Nonmuscle cells are able to regulate CaD activity at several levels in responses to either external or internal signals for controlling a variety of cellular processes such as spreading, shape change, motility, and mitosis. The regulation of CaD activity includes the expression level and the presence of Ca2þ–calmodulin and/or active kinases in a temporally and spatially regulated manner. In the following section, we will discuss how these regulations allow l-CaD plays its roles in nonmuscle cells. 5.2.1. CaD-mediated stabilization of actin microfilament bundles As mentioned above, pituitary cells in response to glucocorticoid express increased amounts of CaD at both mRNA and protein levels, induce actin reorganization to form more stable stress fibers and then inhibit the hormone release (Castellino et al., 1992, 1995). This glucocorticoid-induced actin reorganization and increased CaD can be specifically inhibited by a CaD antisense oligonucleotide in a dose-dependent fashion (Castellino et al., 1995), suggesting that CaD level is an important determinant in controlling hormone release. On the other hand, in many transformed cells, the decrease of l-CaD is observed, concomitant with the loss of stress fibers and reorganization of focal adhesions into podosomes (Nakamura et al., 1993; Novy et al., 1991; Owada et al., 1984; Tanaka et al., 1993). However, in this case, the expression levels of several other actin-binding proteins also change upon transformation, making it difficult to discern the role of CaD from that of other cytoskeletal proteins. Over-expression of wild type or mutant CaD in nonmuscle cells through cDNA transfection or protein microinjection provides a convenient way to evaluate the roles of CaD and its domains in these cells. Cultured nonmuscle cells do not distinguish h- and l-CaD isoforms and can incorporate them equally well into actin filaments at stress fibers and membrane ruffles (Surgucheva and Bryan, 1995; Yamakita et al., 1990). The introduction of l-CaD at a concentration comparable to the endogenous CaD does not affect the actin cytoskeleton (Yamakita et al., 1990), whereas stable over-expression of h-CaD at 4–8 times of endogenous l-CaD in mouse fibroblasts by DNA transfection enhances cell spreading and alters cell morphology (Surgucheva and Bryan, 1995). Furthermore, transient over-expression of l-CaD in nonmuscle cells by DNA transfection inhibits

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the focal adhesion formation and the cell contractility (Helfman et al., 1999). The inhibition requires over-expression of l-CaD that contains intact myosin- and actin-binding domains. These results suggest a role of l-CaD in the regulation of actomyosin contractility and adhesion formation. The apparent discrepancy between protein microinjection and DNA transfections appears to be due to the difference in the amounts of exogenous CaD expressed. The higher exogenous CaD expresses, the more sever disruptions of their stress fibers and focal adhesions are. This point is carefully confirmed by Eves et al in a smooth muscle cell line A7r5 with overexpression of EGFP-tagged l-CaD (Eves et al., 2006). Using an inducible system to express a lower level of CaD in nonmuscle cells, it also confirms the microinjection results that exogenous CaD is mostly associated with microfilaments at stress fibers and membrane ruffles (Yamashiro et al., 2001). Furthermore, it is shown that exogenous l-CaD stabilizes overall actin filaments (Yamashiro et al., 2001). Another approach is to generate stably transfected cell lines overexpressing different amounts of CaD or its mutants and then analyze expression-related phenotype changes in these lines. The extent of a given phenotype observed should parallel the expression level of exogenous proteins. With this criterion, one can rule out the possibility that a given phenotype detected is caused by the disruption of unrelated gene due to the random insertion of plasmid DNA into cell chromosome in DNA transfection experiment. We have over-expressed the C-terminal half (CaD39, aa #244–538) of human fibroblast WI-38 l-CaD II in CHO cells, isolated and characterized stable transfectants which express varying amounts (16–80 times of endogenous CHO CaD) of CaD39 (Warren et al., 1994). Over-expression of CaD39 stabilizes actin microfilament bundles through a reduction in tropomyosin turnover rate. These results support the notion that CaD potentiates nonmuscle tropomyosin binding to actin filaments, and together then stabilizing actin filaments (Warren et al., 1994). As a result of this stabilization, CaD39-expressing cells enhance early cell attachment and spreading in an expression level-dependent manner (Warren et al., 1996). In contrast, none of these effects are observed in the N-terminal half (CaD40)- or full-length CaD-expressing cells. In interphase cells, the expressed CaD39 are found associated with tropomyosin-enriched microfilaments. Like endogenous CaD, the majority of the CaD39 is also modified and released from the microfilaments during mitosis. However, a significant portion of CaD39 undergoes only partial modification and retains in the microfilaments during mitosis. The incomplete release of CaD39 from mitotic microfilaments may be responsible for the increase in the frequency of multinuclear cells in CaD39-expressing cells (Warren et al., 1996), suggesting that the modifications of CaD at mitosis play a critical role for normal cell division, particularly for cytokinesis. We shall discuss the modifications by Ca2þ–calmodulin and by phosphorylation in the next two sections.

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5.2.2. Regulation of CaD activity by Ca2þ–calmodulin in nonmuscle cells Several lines of evidence support the notion that CaD is regulated by Ca2þ– calmodulin in nonmuscle cells. As predicted from the in vitro studies, the binding of Ca2þ–calmodulin to CaD in vivo would reduce its actin-binding affinity and reverse its inhibition of actomyosin ATPase activity. This notion has been first supported by microinjection studies (Yamakita et al., 1990). While the microinjected CaD quickly and faithfully assembles into stress fibers and membrane ruffles of nonmuscle cells, the coinjected calmodulin with CaD appears to retard the incorporation of CaD into these actin microfilament structures. It has been further shown that calmodulin activation by elevated cytoplasmic Ca2þ levels in human fibroblasts blocks CaD inhibitory activity on focal adhesion formation but does not cause CaD to completely dissociate from stress fibers (Helfman et al., 1999). This is in agreement with the in vitro study in which Ca2þ–calmodulin relieves the inhibition of actomyosin ATPase activity while CaD–Ca2þ–calmodulin complex remains bound to the actin–tropomysoin filaments (Pritchard and Marston, 1989). It is thus that a local decrease in affinity of CaD–Ca2þ– calmodulin complex for actin could cause a positional change of CaD that reverses the inhibition of actomyosin ATPase activity (Morgan and Gangopadhyay, 2001). This speculation has been conclusively supported by studies in CHO cells over-expressing CaD mutant defective in Ca2þ– calmodulin binding (Eppinga et al., 2006b; Li et al., 2004). Two well conserved Ca2þ–calmodulin-binding domains have been identified within the C-terminal half (such as CaD39) of CaD, namely sites A and B (Table 1.1 and Fig. 1.2) (Czurylo, 2000; Zhan et al., 1991). Structural studies of CaD–Ca2þ–calmodulin complex suggest that the tryptophan (W) residues within these two sites are crucial for Ca2þ–calmodulin binding (Graether et al., 1997). To study the regulation of CaD by Ca2þ–calmodulin in vivo, a CaD39 mutant (CaD39-AB), in which both tryptophan (W) residues are replaced with alanine (A), is generated and characterized (Li et al., 2004). In vitro, replacing W residues with A residues almost entirely abolishes the binding of CaD39-AB to Ca2þ–calmodulin. CaD39-AB retains actin-binding properties similar to that of wild type CaD39 and is still able to potentiate tropomyosin binding to actin filaments and to inhibit actomyosin ATPase activity. Over-expressed CaD39-AB is incorporated into the actin filaments, but prevents the actin filaments from further bundling into centrally located stress fibers. The disruptions of both stress fibers and focal adhesions in the CaD39-AB-expressing cells cause defects in cell morphology and growth, as well as impair cell motility during wound healing. Cells expressing CaD39-AB migrate much slowly than control cells and change their direction of movement more frequently than control cells (Li et al., 2004). It appears that the functional Ca2þ–calmodulin-binding sites

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on l-CaD are not required for actin-binding but are essential for the dynamics of centrally located stress fibers and their focal adhesions, for the lamellipodia extension, and for the persistence of cell movement (Fig. 1.3). To further study the role of CaD in Ca2þ–calmodulin regulation of cell division, the mitotic behaviors of CaD39-AB-expressing cells are investigated (Eppinga et al., 2006b). Expression of CaD39-AB leads to formation of numerous blebs at early mitosis and distortion of the cleavage furrow during cytokinesis, suggesting that regulation of CaD activity by Ca2þ– calmodulin is important in maintaining proper cortical tensions (Fig. 1.3) (Eppinga et al., 2006b; Li, 2004). During cell division, CaD39-AB becomes concentrated at the membrane cortex region, colocalizing with the cortical actin network (Eppinga et al., 2006b; Li, 2004). This distribution is in contrast to that of the endogenous CaD, which is associated with the stress fiber structures at interphase but becomes diffusely distributed throughout the cytoplasm during cell division. Therefore, Ca2þ–calmodulin regulation of CaD activity is important for the mitosis-specific dissociation of CaD Ca2+/calmodulin ERK

PAK

Cdc2 kinase and ERK

Caldesmon

Reorganization of actin at leading edge

Speed of cell movement, spreading

Lamella extension

Dynamics of stress fibers

Cortical actin rearrangement

Persistence of cell movement

Proper cortical tension

Contraction at contractile ring

Efficient cytokinesis

Figure 1.3 CaD, a downstream effector for multiple signaling pathways, involved in controlling lamellipodia extension, cell motility, cortical tension, and cytokinesis in nonmuscle cells. The binding of Ca2þ–calmodulin and/or phosphorylation of CaD by Cdc2 kinase, PAK, and ERK regulate CaD activity for controlling cell motility and actin cytoskeleton remodeling. The regulations of CaD by different (color-coded) signaling molecules have distinct yet overlapping roles in regulating reorganization of actin at leading edge, dynamics of stress fibers, cortical actin rearrangement, and contraction at contractile ring (indicated by respectively color-coded arrows). Defects in these regulations further reveal various observed phenotypes, which have been supported by the studies of CaD mutants defective in each binding site for these signaling molecules.

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from the cortical actin network. In wild type l-CaD or CaD39-expressing cells, membrane blebs are rare, especially during cell elongation in anaphase. In contrast, greater than half of the cells expressing CaD39-AB exhibit membrane blebs during anaphase (Eppinga et al., 2006b). In CaD39-ABexpressing cells, both actin and CaD39-AB are concentrated at membrane blebs, suggesting that regulation of CaD by Ca2þ–calmodulin may be involved in suppressing bleb formation and maintaining proper cortical tension during cell division (Fig. 1.3). Consistent with the observation of interphase cells in which CaD39-AB does not incorporate into actin bundles, CaD39-AB is excluded from the contractile ring during cytokinesis (Eppinga et al., 2006b; Li, 2004). The distribution of CaD39-AB during cytokinesis is also different from that of another CaD39 mutant (CaD39–6F) defective in all six phosphorylation sites for Cdc2 kinase (Eppinga et al., 2006b; Li, 2004). Interestingly, CaD39–6F-expressing cells also produce membrane blebs during cell division (Eppinga et al., 2006b; Li et al., 2003b), further supporting a role of CaD regulation in maintaining proper cortical tension. We shall discuss the CaD39–6F-expressing cells during division in Section 5.2.3.1. In smooth muscle A7r5 cell line, the endogenous l-CaD is normally localized to stress fibers and membrane ruffles. When these cells are exposed to phorbol ester to induce actin cytoskeleton rearrangements, l-CaD and cortactin are rapidly recruited to the actin core of induced podosomes (Eves et al., 2006; Webb et al., 2006a). The over-expression of l-CaD or CaD39 with intact Ca2þ–calmodulin sites represses the formation of phorbol ester-induced podosomes (Eves et al., 2006). Conversely, siRNA interference of CaD expression increases the formation and dynamics of podosomes (Eves et al., 2006). Furthermore, CaD or CaD39 mutant defective in Ca2þ–calmodulin sites fails to be recruited to podosomes and fails to suppress the podosome formation either in phorbol ester-treated A7r5 cells (Eves et al., 2006). These results suggest that Ca2þ–calmodulin modulates the action of CaD in podosome formation. In vitro CaD inhibits the Arp2/3 complex-mediated actin polymerization (Morita et al., 2007; Yamakita et al., 2003), and directly interacts with cortactin (Huang et al., 2006). Similar to CaD, the Arp2/3 complex and cortactin are also localized to the actin cores of podosomes (Linder and Aepfelbacher, 2003; Webb et al., 2006a). Therefore, the inhibition of podosome formation by CaD is likely through the competition with Arp2/3 complex and/or cortactin for actin binding. This inhibitory role for CaD appears to require functional Ca2þ–calmodulin-binding sites. It further suggests that CaD can act as a repressor for cancer cell invasion by reducing podosome formation. Indeed, over-expression of CaD in Rous sarcoma virus-transformed cells reduces the number of podosomes, decreases the extracellular matrix degradation activity, and suppressing cell invasion and migration (Yoshio et al., 2007). Therefore, CaD may be a

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potential therapeutic tool in cancer cell metastasis. In addition to Ca2þ– calmodulin, the regulation of CaD activity by PAK or ERK phosphorylation can also modulate the podosome formation and dynamics (Gu et al., 2007; Morita et al., 2007). 5.2.3. Regulation of CaD activity by phosphorylation in nonmuscle cells In vitro CaD can be phosphorylated by various kinases listed in Table 1.2, however, only phosphorylation of l-CaD by Cdc2 kinase, ERK, PAK, or v-erbB tyrosine kinase has been observed in nonmuscle cells. Since the significance of tyrosine phosphorylation of CaD by v-erbB tyrosine kinase has been discussed (see Section 3.2 and Table 1.2), we will summarize the physiological significance of CaD phosphorylation by Cdc2 kinase, ERK, and PAK in the following sections. 5.2.3.1. Phosphorylation of l-CaD by Cdc2 kinase Several lines of evidence suggest that mitosis-specific Cdc2 kinase (B-cyclin/Cdk1) regulates l-CaD activity in vivo. CaD is associated with actin stress fibers during interphase, but it is released from actin filaments during mitosis. The phosphorylation of CaD by Cdc2 kinase is shown to cause l-CaD to dissociate from actin filaments during prometaphase (Yamashiro et al., 1990, 1991). CaD is diffusely localized throughout cytoplasm in metaphase. During early stages of cytokinesis, CaD remains diffusely distributed and not associated with the cleavage furrow (contractile ring). At later stages of cytokinesis, some CaD are observed to concentrate in membrane cortex and cleavage furrow (Hosoya et al., 1993). Thus, dissociation of CaD from actin filaments induced by Cdc2 kinase phosphorylation may be required for the assembly and/or activation of contractile ring during cell division (Huber, 1997). To address whether phosphorylation of CaD could regulate organization and contraction of actomyosin during mitosis, CaD mutants defective in all Cdc2 kinase phosphorylation sites are generated and introduced into nonmuscle cells by protein microinjection or DNA transfection experiment. As shown in Table 1.2, rat fibroblast l-CaD contains 7 Cdc2 kinase phosphorylation sites, whereas human fibroblast l-CaD has 6 sites. Microinjection of rat l-CaD mutant defective in all 7 sites (CaD 7th mutant) into Xenopus eggs or CHO cells results in a significant delay of M phase entry (Yamashiro et al., 2001). Similarly, over-expression of CaD39 mutant lacking all 6 sites (CaD39–6F) increases the time required for cells to progress from the start of anaphase to 50% cytokinesis (Eppinga et al., 2006b; Li et al., 2003b). Force-expressed CaD39–6F remains associated with tropomyosin-enriched microfilaments and is concentrated at the contractile ring during cytokinesis, suggesting that phosphorylation of CaD by Cdc2 kinase is required for CaD dissociation from the actomyosin bundles at the equatorial cleavage furrow (Li et al., 2003b). Continuous association

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of CaD39–6F with the contractile ring may interfere with the generation of contractile forces at the equator. Using computer-assisted DIAS programs (Soll and Voss, 1998), detailed motile behaviors of the CaD39–6Fexpressing cells have been analyzed. CaD39–6F-expressing cells exhibit morphological defects during cell division, including an increase in cell size and formation of abnormal blebs. Similar bleb formation is also found in the cells over-expressing human l-CaD defective in all Cdc2 phosphorylation sites (CaD-6F), suggesting Cdc2 phosphorylation of CaD plays a role in regulating cortical tension and inhibiting bleb formation during cytokinesis (Fig. 1.3). However, the cell size of the CaD-6F-expressing cells is indistinguishable from that of the nonexpressing control cells. In contrast, cells expressing the wild type C-terminal fragment (CaD39) show a similar increase in cell size as CaD39–6F-expressing cells. These observations indicate that the N-terminal fragment of CaD may be to alleviate cell size defects caused by the mutations of Cdc2 phosphorylation sites at the C-terminal fragment. It has been proposed that CaD plays dual roles in regulating actin-myosin interactions (Wang, 2001): the C-terminal fragment of CaD inhibits actomyosin ATPase activity; meanwhile, the N-terminal fragment can tether myosin to the vicinity of actin filaments and therefore can facilitate the progression of crossbridge cycles once the inhibitory C-terminal fragment is removed. Although CaD-6F remains associated with actin filaments during cell division due to the loss of Cdc2 phosphorylation, its activity may still be regulated by Ca2þ–calmodulin so that its inhibitory effect on myosin ATPase can be partially reversed. In this scenario, the ability of the N-terminal fragment of CaD-6F to tether myosin would be critical in generating sufficient contractile forces that can contract the mitotic cell into a more compact, normal shape and size. The phenotypic differences between the expressions of the C-terminal half and the full-length proteins are also observed in CaD- and CaD39-expressing cells. In contrast to CaD39, over-expression of full-length CaD cannot stabilize tropomyosin in interphase (Li, 2004; Warren et al., 1994), suggesting that the effects of the full-length and the C-terminal fragment of CaD on actin dynamics are not equivalent. The blebs observed in dividing CaD39–6F-expressing cells are similar to that described early for dividing CaD39-AB expressing cells but different from those previously reported in the cells expressing chimeric tropomyosin isoform hTM5/3 (Wong et al., 2000). Compared to the small, transient, and numerous blebs in both CaD39–6F and CaD39-AB-expressing cells, the blebs formed in the hTM5/3-expressing cells are large protracted membrane bulges. Blebs are a dissociation of the membrane from the underlying cytoskeleton and bulges are deformations of the cytoskeleton itself (Charras et al., 2005; Eppinga et al., 2006b). The difference in cortical tension between cells that bleb and cells that bulge is reflected in their response to hypotonic solution. During mitosis, CaD39–6F- and CaD39-AB-expressing

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cells show less resistance to swelling in hypotonic media, whereas hTM5/3expressing cells show more resistance (Eppinga et al., 2006b). Ca2þ–calmodulin and Cdc2 kinase control CaD activity spatially during cytokinesis to promote cell division. Force-expressed CaD39–6F localizes to the contractile rings and the membrane cortexes, which is very similar to the distribution of Ca2þ–calmodulin in dividing cells (Li et al., 1999). It is possible that Ca2þ– calmodulin regulation can partially relieve the inhibitory effects of CaD39–6F on contractile rings, allowing constriction at the equator but requiring more time. Similar to CaD39–6F, force-expressed CaD39-AB mutant protein is also enriched in the membrane cortex (Eppinga et al., 2006b; Li, 2004; Li et al., 2003b), at where both mutant proteins could cause local disruption of membrane tension, leading to the production of blebs. In summary, Cdc2 kinase together with Ca2þ–calmodulin dynamically control CaD inhibition of tropomyosin-actin activated myosin ATPase to regulate division speed and to suppress membrane blebs (Fig. 1.3). In prostate cancer cells, the up-regulation of Cdc2 message and protein via the expression of avb3 integrin has been shown to promote cell migration without affecting cell adhesion to integrin ligands (Manes et al., 2003). A correlation between Cdc2 levels and more migratory phenotypes on integrin ligands is also observed in other human cell lines, including fibrosarcoma cell line HT1080 and HeLa. Cdc2 has been shown to specifically associate with cyclin B2 and the activated B2-cyclin/Cdk1 kinase not only colocalizes with CaD to the membrane ruffles of motile cells but also phosphorylates CaD. As a result, this up-regulation of Cdc2 in response to the expression of avb3 integrin greatly enhances cell migration. Functional Cdc2 kinase phosphorylation sites on CaD are required for enhancing cell migration. Together, these results indicate that B2-cyclin/Cdk1 kinase is a downstream effector of the avb3 integrin, regulating CaD activity to promote cell migration. Recent studies on osteoclast biology and cancer cell metastasis further support this notion. In osteoclasts, the avb3 integrin is known to organize the actin ring (cloud) surrounding the actin core (column) of podosomes, which are essential for extracellular matrix degradation, invasion, and migration in vivo (Chabadel et al., 2007; Linder, 2007; Saltel et al., 2008). The l-CaD found in the actin cores of podosomes is able to regulate the formation of podosomes (Eves et al., 2006). Furthermore, the expression of tumor-specific avb3 integrin has been shown to promote metastasis of prostate and breast cancer cells to bone (McCabe et al., 2007; Sloan et al., 2006; Takayama et al., 2005). K-cyclin (a D-like cyclin) encoded by Kaposi sarcoma-associated herpes virus is able to activate host cell Cdk6 during infection. Using a kinase substrate tracking and elucidation screen (Knebel et al., 2001), CaD has been identified as one of novel substrates for K-cyclin/Cdk6 kinase (Cuomo et al., 2005). The identified phosphorylation sites on human h-CaD are T730, T753, S759, and S789, which are identical to those sites phosphorylated

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by Cdc2 kinase (B-cyclin/Cdk1) during cell division. Similar to Cdc2 kinase phosphorylation of CaD (Mak et al., 1991), phospho-CaD by K-cyclin/Cdk6 drastically reduces its binding ability to actin filaments and to Ca2þ–calmodulin, and the preincubation with actin filaments or Ca2þ– calmodulin also decreases its phosphorylation activity by about 50% (Cuomo et al., 2005). In vivo, expression of K-cyclin through Cdk6-dependent phosphorylation of l-CaD affects actin cytoskeleton organization, leading to changes in cell shape, loss of focal adhesions, and increased number of membrane protrusions (Cuomo et al., 2005). These results indicate that K-cyclin contributes to the viral oncogenecity through the activation of host Cdk6, phosphorylation of CaD, and then rearrangement of actin cytoskeleton for shape changes. 5.2.3.2. Phosphorylation of l-CaD by ERK ERK, another proline-directed protein kinase, is known to recognize same (S/T)-P-X-X consensus sequence as Cdc2 kinase, but preferentially phosphorylates S759 and S789 on human h-CaD, S504 and S534 on human l-CaD, S717 on chicken h-CaD, or S497 and S527 on rat l-CaD (Table 1.2) (Adam et al., 1992; Childs et al., 1992; D’Angelo et al., 1999). Phosphorylation of CaD by Cdc2 kinase and ERK appears to affect differently the structure and function of CaD. In vitro, ERK-mediated phosphorylation of CaD has little effect on the CaD-Ca2þ– calmodulin interaction (Adam et al., 1992; Childs et al., 1992), whereas Cdc2-mediated phosphorylation of CaD at 6 or 7 sites reduces the affinity of CaD to Ca2þ–calmodulin (Yamashiro et al., 1991, 1995). Furthermore, phosphorylation of CaD by ERK at sites near the C-terminus has a relatively small effect on the CaD and actin interaction (Foster et al., 2004; Huang et al., 2003), but phosphorylation by Cdc2 kinase drastically reduces the binding of CaD to actin filaments. Thus, it is important to apply specific kinase inhibitor in vivo to identify which kinase causes a CaD phosphorylation-dependent effect, in addition to showing the activation of specific kinase. One such example is described above for B2-cyclin/Cdk1 involved in the avb3 integrin-mediated cell migration in osteoclasts and certain cancer cell metastasis (Manes et al., 2003). There are few reports investigating the role of l-CaD phosphorylation by ERK in actin cytoskeleton remodeling and motility in nonmuscle cells. With the availability of ERK phosphorylation site-specific antibodies (anti-pCaD759 and anti-pCaD789) (D’Angelo et al., 1999), it is shown that l-CaD is rapidly phosphorylated only at S789 (equivalent to S534 of human l-CaD) as cultured smooth muscle cell line progresses from serumstarved conditions into exponential growth conditions. This phosphorylation is markedly inhibited by the MEK specific inhibitor, suggesting that ERK but not Cdc2 kinase is responsible for this in vivo phosphorylation (D’Angelo et al., 1999). An ERK phosphorylation-dependent conformational change has been detected by photo-crosslinking and 3D reconstruction of thin

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filaments (Foster et al., 2004). This conformational change provides a mechanism to relieve the CaD inhibition of actomyosin ATPase activity without leaving of CaD from the actin filaments (Fig. 1.4). In nonmuscle cells, the level of l-CaD phosphorylation at S504 and S534 undergoes dynamic changes during cell cycle (Kordowska et al., 2006a). The spatial and temporal distributions of phosphorylated CaD appears to correlate with the level of stress fibers in a reciprocal manner. These results suggest that phosphorylation of these residues of l-CaD facilitates stress fiber disassembly and then actin cytoskeleton remodeling (Kordowska et al., 2006a) (Fig. 1.4). Unfortunately, this study did not attempt to distinguish whether ERK or Cdc2 kinase phosphorylates l-CaD.

*C *R a 2+-c by ever alm PA sib odu l K or e ph lin ER osp K ho

ryl

ati

Myosin + actin·TM·CaD Inhibitory state

Cytoskeleton dynamics and cell motility

ulin od lm c2 + -ca y cd b a2 t C tion a an nd oryl h bu *A osp h *P

on

• Enhances TM binding to actin • Inhibits actin bundling • Inhibits myosin ATPase activity

Activated state

Released state

Myosin + actin·TM·CaD

Myosin + actin·TM + CaD

• Facilitates assembly of stress fibers • Stabilizes structures of stress fibers and focal adhesions

• Allows the rearrangement of filamentous structure • Activates myosin ATPase activity

Figure 1.4 A proposed three-state model for the regulations of CaD activity in modulating cytoskeleton dynamics and cell motility. In nonmuscle cells, CaD can exist in three states. CaD associated with tropomyosin (TM)-containing actin in the absence of any regulators adopts an inhibitory conformation (in red), which stabilizes actin filaments while preventing further assembly into bundles and inhibiting actomyosin ATPase activity. The presence of moderate Ca2þ–calmodulin in interphase cells changes the conformation of CaD into an activated state (in green) and allows the formation of stress fibers. Excessive Ca2þ–calmodulin during cell division, as well as phosphorylation of CaD at interphase or at mitosis, locks CaD in a released state (in purple) in which CaD becomes dissociated (or partial dissociated in case of PAK or ERK) from actin filaments and the effects of CaD on actin reorganization and actin– myosin interaction are reversed. Actin cytoskeleton dynamics and cell motility are determined by the balance among these three states.

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5.2.3.3. Phosphorylation of l-CaD by PAK PAK is emerging as a major regulator of CaD-mediated actin dynamics in vivo. The PAK is a serine/ threonine protein kinase whose activity is stimulated by the binding of active Rac and Cdc42 GTPases. Accumulated lines of evidence suggest that PAK acts as downstream effectors of Rac and Cdc42 GTPases to regulate a range of biological activities including the control of cytoskeletal dynamics (Bishop and Hall, 2000). When stimulated by PDGF or during wound healing, PAK redistributes from the cytosol to the lamellipodia at the leading edge of a polarized cell (Dharmawardhane et al., 1997). Expression of kinase-active PAK mutant induces large, polarized lamellipodia at the leading edge of translocating 3T3 fibroblasts and caused increased cell motility (Sells et al., 1997, 1999). On the other hand, expression of kinasedead PAK mutant in a breast cancer cell line blocks the formation of ruffles and filopodia, thus inhibiting cell transmigration across a porous membrane (Adam et al., 2000). All these observations indicate that the kinase activity of PAK is important for membrane extension at the leading edge of a migrating cell. In vitro biochemical assays have identified a number of kinase substrates, including CaD, on the cytoskeleton system that can account for some, if not all, of the PAK effects on membrane extension and cell motility (Sander et al., 1998; Van Eyk et al., 1998). The phosphorylation sites for PAK have been mapped to the two serine residues at S458 and S489 of human fibroblast l-CaD (Foster et al., 2000). While PAK-phosphorylation leads to only a moderate reduction in its binding affinity for actin filaments in vitro, it significantly reduces the ability of CaD to inhibit actomyosin ATPase activity (Foster et al., 2000). Using antibody specific to PAK phosphorylation sites on CaD (McFawn et al., 2003), it has been shown that the putative PAK sites are phosphorylated in vivo and that this phosphorylation is increased when CHO cells are stimulated to migrate in wound healing assays (Eppinga et al., 2006a). To study the phosphorylation of CaD by PAK, and to determine the role of this phosphorylation in regulating cytoskeleton dynamics, we generate two site-directed mutants of CaD39 with opposite phosphorylation properties: one of them, CaD39-PAKA, bears mutations of the serine residues into alanines so that it cannot be phosphorylated by PAK; the other one, CaD39-PAKE, mimics the phosphorylation state of CaD39 by replacing the two serine residues with glutamic acids. In vitro, nonphosphorylatable CaD39-PAKA has similar binding properties and similar inhibition of actomyosin ATPase activity as wild type CaD39, whereas CaD39-PAKE exhibits reduced affinity for both Ca2þ–calmodulin and actin as well as reduced inhibition of actomyosin ATPase activity in a similar manner as PAK-phosphorylated CaD39 (Eppinga et al., 2006a). Despite their differences in in vitro properties, both CaD39-PAKA and CaD39PAKE are able to incorporate into stress fibers when stably expressed in CHO cells. Expression of CaD39-PAKA enhances cell spreading and

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protects stress fibers from cytochalasin depolymerization but not to the extent of that by CaD39 expression. In contrast, CaD39-PAKE-expressing cells fail to protect stress fibers from depolymerization and fail to enhance cell spreading. However, Both CaD39-PAKA- and CaD39-PAKE-expressing cells are less polarized and, during wound healing, exhibit defects in lamellipodia extension and migrate slower than the control cells (Eppinga et al., 2006a). Neither the phosphorylated nor unphosphorylated state is sufficient to allow dynamic rearrangements of actin cytoskeleton required for normal cell motility. Thus, PAK-phosphorylation of l-CaD is a dynamic process involved in the regulation of stress fiber integrity, cell motility, and membrane extension at the leading edge of translocating cells (Fig. 1.3). To examine whether the PAK phosphorylation of l-CaD is also necessary during cytokinesis, mitotic cells expressing CaD39-PAKA and CaD39PAKE have been analyzed by DIAS program (Soll and Voss, 1998; Wong et al., 2000). CaD39 mutants that mimic either the phosphorylated state (CaD39-PAKE) or nonphosphorylated state (CaD39-PAKA) have no statistical effect on cell division speed compared with wild type cells. However, CaD39-PAKE speeds division significantly faster than cells expressing opposite mutant CaD39-PAKA, indicating that PAK may influence CaD regulation of cytokinesis in vivo, but if it does, this role is minor compared with the Ca2þ–calmodulin and Cdc2 kinase (Eppinga et al., 2006b).

6. CaD, Podosomes, and Diseases Podosomes are highly dynamic cell-adhesion structures that degrade the extracellular matrix by secreting matrix metalloproteases, allowing cells to cross the basal lamina and to invade tissues in physiological and pathological conditions (Buccione et al., 2004; Linder, 2007). The l-CaD has been shown to be not only an integral component (in the actin core) of podosomes but also an important regulator of podosome formation and dynamics (Eves et al., 2006; Gu et al., 2007; Morita et al., 2007; Tanaka et al., 1993). Ectopic expression of human l-CaD in RSV-transformed fibroblasts is able to reduce the number of podosomes formed, to decrease the extracellular matrix degradation activity, and suppress the cell invasion. Conversely, down-regulation of CaD expression by siRNA treatment enhances the formation of podosomes and cell invasion (Yoshio et al., 2007). The molecular mechanism for the CaD action in podosome formation and dynamics remains unclear. Studies from osteoclasts and macrophages reveal that the podosome core is formed by the activation of transmembrane receptor CD44 by ligand (hyaluronan) binding and subsequently by the stabilization of a signaling system for actin polymerization and organization through the direct binding of CD44 to N-WASP (neuronal WASP)

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(Bourguignon et al., 2007). The components of this signaling system include N-WASP, WIP (WASP interacting protein), Arp2/3, cortactin, and formin (Saltel et al., 2008). The activated N-WASP is known to enhance the Arp2/3 complex for branched actin polymerization (Linder et al., 1999). Through its SH3 domain, cortactin can bind and further activate N-WASP for enhancing N-WASP-mediated Arp2/3 activation (Ammer and Weed, 2008; Webb et al., 2006a). This activation can be negatively or positively regulated by phosphorylation at the helical, proline-rich (HP) region of cortactin with Src tyrosine kinase or ERK, respectively (Martinez-Quiles et al., 2004). Translocation of additional cortactin to the actin core of podosome requires its 4th actin-binding repeat (ABR) region (Webb et al., 2006a). Furthermore, cortactin binds the Arp2/3 through its N-terminal acidic (NTA) domain, resulting in direct activation of the Arp2/3 nucleation activity. Through these multiple mechanisms, cortactin stabilizes the branched actin networks and enhances podosome formation (Webb et al., 2006a). Phosphorylation of cortactin by PAK at the first ABR (Webb et al., 2006b) and by ERK at HP (Campbell et al., 1999; MartinezQuiles et al., 2004) could further enhance actin polymerization and branched actin network formation (Ammer and Weed, 2008). In contrast, CaD found in the podosmoe core can inhibit the Arp2/3 complexmediated actin polymerization and thereby inhibits podosome formation (Morita et al., 2007; Yamakita et al., 2003). Phosphorylation of CaD by Cdc42-activated PAK further increases its ability to compete with Arp2/3 complex and enhances its inhibitory effect on podosome formation (Morita et al., 2007). On the other hand, active Cdc42 has been shown to directly bind N-WASP, activate Arp2/3 complex, and promote podosome formation in vascular endothelial cells (Moreau et al., 2003, 2006). Therefore, changes in the balance between the CaD activity regulated by PAK and the N-WASP-Arp2/3-cortactin pathway determine the podosome assembly/ disassembly (Morita et al., 2007). As discussed above, cortactin is an important regulator in stabilizing branched actin network for podosome formation (Ammer and Weed, 2008). The recent finding that CaD directly interacts with cortactin (Huang et al., 2006) may suggest another mechanism for CaD to regulate podosome formation. Both cortactin and CaD are also phosphorylated by ERK. Whereas phosphorylation of cortactin by ERK is shown to enhance the podosome formation (Martinez-Quiles et al., 2004), ERK phosphorylation of CaD appears to regulate the size and life time of podosomes (Gu et al., 2007). The phorbol ester-treated vascular smooth muscle A7r5 cells produce numerous podosomes (Eves et al., 2006; Gu et al., 2007). Overexpression of wild type l-CaD or mutant l-CaD defective in ERK phosphorylation sites decreases the average number of podosomes per cell in phorbol ester-treated A7r5 cells (Gu et al., 2007). However, the podosomes formed in mutant CaD-expressing cells are smaller in size and shorter in life

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time as compared to that observed in wild type CaD-expressing cells suggesting that CaD activity regulated by ERK may modulate the podosome dynamics, size, and turn over time (Gu et al., 2007). Therefore, it is likely that CaD through its phosphorylation by at least PAK and ERK can effectively modulate the podosome formation and dynamics. It remains to be determined whether CaD activity also plays a role in further modulating the secretion of matrix metalloproteases from podosomes. Bladder outlet obstruction is a common medical problem, especially in men with benign prostatic hyperplasia. This obstruction is known to induce numerous morphological and functional changes in detrusor smooth muscle of the bladder wall, leading to progressive bladder dysfunction (impairment of the ability of the bladder to store and to empty urine) (Levin et al., 2000). In the rabbit model of partial bladder outlet obstruction (PBOO), detrusor smooth muscle first undergoes hypertrophy to compensate for the increased force required to expel urine against the obstruction (compensated state), and progresses to bladder dysfunction (decompensated state) (Chacko et al., 2004). The detrusors from normal bladder express predominantly h-CaD and very little of l-CaD. During the 8-week course of obstruction, the expression of h-CaD increases significantly, reaches a plateau level at 2 weeks, and then drops but still maintains slightly higher than the normal control (Yang et al., 2008). In addition, obstructed detrusor smooth muscles continuously express increased amounts of l-CaD. By 4 and 8 weeks of PBOO, the increases in l-CaD expression are almost 8- and 25-folds, respectively, higher than the control. Another investigator has used the same rabbit PBOO model with different regime, in which obstructed animals are divided into compensated and decompensated groups. Basically, similar conclusion on the obstruction-induced CaD isoform changes has been obtained (Zhang et al., 2004). Therefore, the level of l-CaD together with the ratio of h-CaD/l-CaD may use as markers to monitor the degree of detrusor smooth muscle remodeling and dysfunction in PBOO. Similar to PBOO, detrusor smooth muscle of bladder wall also undergoes hypertrophy in the rabbits with alloxan-induced diabetes mellitus (Mannikarottu et al., 2005). However, in this model, the up-regulation of thin filament proteins, such as calponin, tropomyosin, and h-CaD may be responsible for the observed decreases in contractile force and sensitivity (Changolkar et al., 2005; Gupta et al., 1996). In human, at least four isoforms of l-CaD have been cloned (Fig. 1.1). HeLa type l-CaD uses different promoter and exon 1b from CALD1 gene to produce slightly smaller CaD proteins than WI-38 fibroblast type l-CaD. Interestingly, using exon 1b-specific antibody, it has been shown that the HeLa l-CaD is abnormally expressed in vascular endothelial cells of many tumor tissues derived from a variety of organs but not found in normal blood vessels (Zheng et al., 2005b). These HeLa l-CaD-positive endothelial cells appear to acquire motile phenotype, lose vinculin-containing focal

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adhesions and form podosome-like structures. The HeLa type l-CaD is also expressed in the cytoplasm of normal epithelial cells and carcinomas (Zheng et al., 2005b). Studies on normal brain microvasculature and glioma microvessels with a combination of laser-capture microdissection, RT-PCR, Western blot, and immunohistochemistry conclude that mis-splicing of exon 1b, resulting in the up-regulated expression of HeLa type l-CaD, is exclusively found in the endothelial cells but not glial cells of glioma microvessels (Zheng et al., 2004). The drastic increase in HeLa type l-CaD expression, concomitant with the down-regulated expression of tight junction proteins in the endothelial cells of patients with glioma (Zheng et al., 2004), may account for the source of l-CaD detected with high sensitivity and specificity in cerebrospinal fluid (Zheng et al., 2003) and serum (Zheng et al., 2005a). Thus, the serum l-CaD level can be a good noninvasive biomarker to detect patients with glioma. Furthermore, abnormally expressed HeLa l-CaD is found to be phosphorylated by Cdc2 kinase and associated with a well-known DNA-binding protein, Ki67, which is only expressed in cell cycle-activated (proliferative) cells but not in quiescent G0 cells. Nuclear localization of HeLa l-CaD is exclusively detected in the Ki67-positive proliferative cells and is independent of other actin binding proteins, such as tropomyosin, vinculin, or gelsolin. In contrast, this Hela l-CaD is found to remain in the cytoplasm of the Ki67-negative cells (Zheng et al., 2007). The nuclear translocation of this HeLa l-CaD potentially suggests a new role for CaD in regulating nuclear functions. Encouraging from the finding that over-expression of l-CaD in cultured fibroblasts results in losses of actin stress fibers, focal adhesions, and myosin II-dependent cell contractility (Helfman et al., 1999), several investigators have attempted to develop therapeutic strategies to use CaD expression for anti-angiogenesis (Numaguchi et al., 2003; Yokouchi et al., 2006), for protecting endothelial cell barrier dysfunction (Haxhinasto et al., 2004), and for lowering intraocular pressure in the eye of glaucoma patients (Gabelt et al., 2006; Grosheva et al., 2006). Table 1.4 briefly summarizes some of examples in developing therapeutic strategy for various diseases.

7. Concluding Remarks CaD is a downstream effector for multiple signaling pathways in smooth muscle and nonmuscle cells, which are involved in the regulations of Ca2þ–calmodulin concentration and various kinase activities. In permeabilized smooth muscle strips, synthetic peptides antagonists encompassing the actin-, Ca2þ–calmodulin- or myosin-binding domain compete with endogenous CaD and induce muscle contraction, suggesting that in vivo CaD plays an inhibitory role. Upon the stimulation of smooth muscle by

Table 1.4 Examples for strategies to use CaD expression as a therapeutic tool for various diseases

Investigators

Cells or tissues

(I) D. Ingber (Numaguchi et al. (2003))

Cultured bovine capillary endothelial cells

(II) S. Chacko (Shukla et al. (2004))

Bladder smooth muscle (BSM) cell line derived from obstructed rabbit bladder smooth muscle

DF-1 cells (an (III) A. B. Moy avian l-CaD(Haxhinasto et al. null cell line) (2004))

Force-expressed CaD and vector used

Purposes

Results

Adenoviral vector encoding GFP-rat l-CaD (AdGFPCaD) under Tetoff control (AdTetoff ) In addition to transfection with pcDNA3.1chicken h-CaD, cells are exposed to intermittent agonist-induced contraction

Can l-CaD expression represent a potential target for antiangiogenesis therapy?

Adenoviral expression system, Ad-human l-CaD and Ad control constructs

Can l-CaD expression protect cell membrane integrity from adenovirus infection in DF-1 cells?

Expression of l-CaD inhibits contractility, disrupts stress fibers and focal adhesions, increases apoptosis but inhibits cell growth. Agonist-induced intermittent contraction increases h-CaD expression and the restoration of h-CaD alters cell morphology and cytoplasmic filament organization. Transfection of Ad control reduces transcellular resistance, via disruption of cell– cell and cell–matrix adhesion. Expression of l-CaD protects cells from reducing transcellular resistance.

Can stable expression of h-CaD and/or agonist-induced contraction in BSM cells restore their differentiated smooth muscle phenotype?

(continued)

46 Table 1.4 (continued) Investigators

Cells or tissues

Force-expressed CaD and vector used

Purposes

Results

In VSMC, l-CaD expression results in the progressive loss of stress fibers and focal adhesions, the increase in apoptosis, and decrease in cell growth, and inhibits PDGFinduced cell migration. In injured carotid artery, l-CaD expression inhibits neointimal formation. In HTM cells, l-CaD expression results in changes in contractility, actin cytoskeleton, and adhesions.

(IV) Y. Numaguchi Cultured human aorta vascular (Yokouchi et al. smooth muscle (2006)) cells (VSMC) and rat right carotid artery after inflicting a balloon injury

Ad-rat l-CaD

Can l-CaD be a cytostatic agent to inhibit neointimal formation after angioplasty by suppressing VSMC growth and migration?

Cultured human (V) B. T. Gabelt trabecular (Gabelt et al. meshwork (2006); Grosheva (HTM) cells and et al. (2006)) organ cultured human and monkey anterior segments of the eye

Ad-rat l-CaD-GFP (l-CaD fused to GFP protein)

Can l-CaD expression lowers intraocular pressure and increases outflow facility from the anterior chamber of the eye? (potential gene therapy for glaucoma)

(VI) K. Sobue (Yoshio et al. (2007))

Rous sarcoma virustransformed 3Y1 cells

pcDNA3.1-HA-CaD (HA-tagged l-CaD)

Can CaD act as a repressor of cancer cell invasion?

In organ cultured anterior segments, l-CaD expression results in increases in the outflow facility, leading to lowering the intraocular pressure. Expression of l-CaD reduces podosome numbers, extracellular matrix degradation activity, and cancer cell invasion.

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various agonists, the regulation of CaD activity can be observed and accompanied with muscle contraction and/or actin cytoskeleton remodeling. However, a generalized correlation between CaD regulation and smooth muscle responses cannot be easily concluded at the present time, due to variations among agonists, smooth muscle tissue types, differentiation states of smooth muscle, and animal species used in the experiments. Figure 1.4 summarizes what we know about the roles of l-CaD regulation by Ca2þ–calmodulin and/or phosphorylation by Cdc2 kinase, ERK, or PAK in cytoskeleton dynamics and cell motility. The regulation of l-CaD by Ca2þ–calmodulin is important in maintaining the integrity of focal adhesion and stress fiber structures. Mutation in Ca2þ–calmodulin-binding sites not only leads to the inability of the mutant to incorporate into stress fibers, but also disrupts the structure of stress fibers itself, which in turn result in the defects in cell morphology, cell cycle progression, and cell motility (Li et al., 2004). This finding seems to be paradox to the classic theory in which the binding of CaD can stabilize the actin stress fibers. CaD can exist in three conformational (inhibitory, activated, and released) states in nonmuscle cells (Fig. 1.4). CaD serves dual functions in regulation of stress fiber assembly. Firstly, CaD in an inhibitory state together with tropomyosin prevents actin filaments from bundling by bundling protein such as fascin. The presence of Ca2þ–calmodulin likely in the ternary structure with actin, tropomyosin and CaD (possessing functional Ca2þ–calmodulin-binding sites), is required to regulate CaD activity so that tropomyosin and CaD are in appropriate position (i.e., an activated state) to allow the binding of fascin to actin filaments, leading to assembly of stress fibers. Once stress fibers are formed, associated CaD (in an activated state) can stabilize the structures and prevent it from disassembly, possibly by inhibiting the activites of actin-severing proteins such as gelsolin, cofilin. Phosphorylated CaD by Cdc2 kinase (in a released state) during mitosis completely dissociates from actin bundles at the contractile ring. Forceexpressed CaD39–6F defective in all Cdc2 phosphorylation sites remains associated with actin filaments at the contractile ring during cytokinesis. However, these cells are still able to progress to anaphase, form cleavage furrows and divide symmetrically, although the total elapsed time for this period is significantly longer (Li et al., 2003b). These results further suggest that CaD-phosphorylation by Cdc2 kinase is not the only way to release the inhibitory effect of CaD on actomyosin ATPase activity and actin cytoskeleton rearrangement during cell division. One of the potential regulators is Ca2þ–calmodulin. It has been reported that Ca2þ-bound calmodulin, an active form, locates at cell cortex region and contractile ring during cell division (Li et al., 1999). The abundant Ca2þ–calmodulin found at the contractile ring may effectively release the inhibitory action of associated CaD39–6F, allowing the constriction of cleavage furrow. In contrast, accumulated Ca2þ–calmodulin at cell cortex region may not be enough to

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compensate the defect generated by CaD39–6F mutant. Therefore, CaD39– 6F-expressing cells produce numerous blebs during cytokinesis (Eppinga et al., 2006b; Li et al., 2003b). Studies with Ca2þ–calmodulin-binding site mutant also show that during cell division, CaD39-AB colocalizes with the actin filaments at the cortex regions but excludes from the contractile rings due to Cdc2 kinase phosphorylation (Eppinga et al., 2006b; Li, 2004). Although CaD39-AB-expressing cells are able to progress through cytokinesis, they also emerge a numerous number of blebs, exhibit distortion at the cleavage furrows, and increase the cell shape complexity (Eppinga et al., 2006b; Li, 2004). During cytokinesis, Ca2þ–calmodulin and Cdc2 kinase cooperatively release the brake through CaD whereas PAK regulation of CaD has only a minor influence. It would be interesting to express in nonmuscle cells a mutant that is deficient in both Ca2þ–calmodulin and Cdc2 kinase regulatory pathways. It is likely that such a mutant would not be released from cortical actin filaments or from the contractile ring, causing a severe delay in cytokinesis. Finally, motility defects observed in CaD39-AB-expressing cells resemble those previously reported in calmodulin-deficient cells (Ranta-Knuuttila et al., 2002; Walker et al., 1998), suggesting that CaD is involved in Ca2þ–calmodulin control of cell migration and plays an important role in this regulatory pathway. In interphase cells, phorphorylation of CaD by PAK or ERK may partially dissociate CaD into an activated state (Fig. 1.4) and release its inhibitory action on actomyosin ATPase activity and actin cytoskeleton rearrangement. PAK-phosphorylation of CaD is a dynamic process (Eppinga et al., 2006a). Both over- and under-phosphorylation cause retardation of membrane extension, supporting the notion that lamellipodia dynamics at the leading edge requires the phosphorylation of CaD to be regulated in a temporally and spatially specific manner. This requirement does make sense because the initial extension of the membrane needs the actin rearrangement and polymerization, which in turn requires the destabilization of existing filaments, yet later on the stabilization of membrane extension requires stabilization of actin and focal adhesions at the leading edge. Although mutations in PAK-phosphorylation sites of CaD do cause defects in cell motility, the defects do not replicate those previously observed in the expression of kinase-active and kinase-dead forms of PAK (Adam et al., 2000; Sells et al., 1997). Therefore, it is likely that PAK possesses multiple downstream targets on the cytoskeleton and its regulations of these targets work synergistically to mediate PAK-regulation of cytoskeleton and cell motility. When compared with the motile behaviors in the CaD39-AB-expressing cells, mutations at PAK phosphorylation sites do not cause a defect in persistence of cell movement (Fig. 1.3), suggesting that integrity of stress fiber structures play an important role in directionality of cell movements. Reversible phosphorylation of CaD by PAK allows the dynamic rearrangement of actin that is required during cell migration (Eppinga et al., 2006a).

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The importance of CaD regulation by PAK during cell migration has been further confirmed by showing that it can control the formation of podosomes in Rous sarcoma virus-transformed cells (Morita et al., 2007). Although the molecular mechanism remains unclear, the PAK-phosphorylated CaD can further inhibit the formation of Arp2/3-induced podosomes. Thus, it appears that deregulation of PAK phosphorylation of CaD can contribute to aggressive metastatic potential of transformed cells. The two PAK phosphorylation sites are located adjacent to the two Ca2þ–calmodulin binding sites. Ca2þ– calmodulin binds CaD39-PAKE less efficiently than CaD39 or CaD39PAKA, yet neither of the CAD39-PAK mutants mimics the CaD39-AB mutant defective in lamelle extension and speed of cell movement (Fig. 1.3), suggesting that PAK phosphorylation of CaD has a distinct physiological role. Analysis of double mutants at the PAK sites and Ca2þ–calmodulin sites could determine whether PAK phosphorylation of CaD can have an additive effect on cytokinesis. ERK phosphorylates CaD at the same two sites by Cdc2 kinase. The role of CaD activity regulated by ERK during cell division may be similar to but difficult to separate from that by Cdc2 kinase. However, in interphase cells, it has been shown that phosphorylation level of CaD by ERK inversely correlates with the level of stress fibers, suggesting that ERK-phosphorylated CaD may inhibit the assembly of stress fibers. ERKphosphorylated CaD colocalizes with nascent focal adhesion during spreading, suggesting ERK phosphorylation of CaD may play important role in cell spreading and migration (Kordowska et al., 2006a). Analysis of double mutants at the PAK sites and ERK sites may further advance our understanding of cell spreading and migration. Stabilization of actin filaments by CaD in vitro does not necessarily means to stabilize stress fibers in vivo. In fact, transient expression of CaD in nonmuscle cells disrupt stress fibers and focal adhesions, and consequently, the cell contractility (Helfman et al., 1999). It is under the regulation of Ca2þ–calmodulin that actin can bundle together and CaD functions to stabilize stress fibers (Li et al., 2004). The regulation of CaD activity by Cdc2 kinase, ERK, or PAK can cause complete or partial dissociation of CaD from actin filaments, allowing the rearrangement of actin filaments and contraction of actomyosin. The regulatory effect of CaD on cytoskeleton and cell motility depends on the balance among three different (inhibitory, released, and activated) states of CaD (Fig. 1.4). The h-CaD is an excellent marker for differentiated smooth muscle. The detection of l-CaD in cerebrospinal fluid and serum may be a useful tool for diagnosis of glioma and monitoring the tumor prognosis during treatment. The expression of abnormal splicing variant of HeLa type l-CaD in tumor neovasculization and its nuclear localization may provide a new marker of angiogenic endothelial cells. The ratio of h-CaD/l-CaD represents smooth muscle differentiation-dedifferentiation states and also

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provides a good marker for the bladder detrusor smooth muscle remodeling and dysfunction. In addition, CaD can be an effective therapeutic tool for various diseases including cancer invasion, anigiogenesis, and glaucoma. Of course, the field of developing therapeutic strategy with CaD is still in its infancy and remains to be further investigated.

ACKNOWLEDGMENTS We would like to thank J. L-C. Lin, J-Y. Choi, W. Pierce, J. Sholley, A. Matveia, and A. Van Winkle for their technical assistance. This work was supported by NIH grants HD18577 and HL76810 to J.J.-C.L. and grant HL086720 to J.-P. J.

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Hypothalamic Neural Systems Controlling the Female Reproductive Life Cycle: Gonadotropin-Releasing Hormone, Glutamate, and GABA Jacqueline A. Maffucci* and Andrea C. Gore*,†,‡ Contents 1. Introduction 2. The HPG Axis 3. The Reproductive Life Cycle in Females 3.1. Development and the attainment of reproductive competence 3.2. Reproductive senescence 3.3. The GnRH-glutamate-GABA connection: An introduction 4. Glutamate and the HPG Axis 4.1. Expression of glutamate receptors in the hypothalamus, and specifically on GnRH cells 4.2. Structure-function properties of glutamate receptors 4.3. The glutamate-GnRH connection across the life cycle 5. Gamma-Aminobutyric Acid (GABA) and the HPG Axis 5.1. Expression of GABA receptors in the hypothalamus and specifically on GnRH cells 5.2. Structure-function properties of GABA receptors 5.3. The GABA-GnRH connection across the life cycle 6. Glutamate/GABA Interactions 7. Conclusion Acknowledgments References

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Abstract The hypothalamic-pituitary-gonadal (HPG) axis undergoes a number of changes throughout the reproductive life cycle that are responsible for the development, puberty, adulthood, and senescence of reproductive systems. This natural * { {

Institute for Neuroscience, University of Texas, Austin, Texas 78712 Division of Pharmacology and Toxicology, University of Texas, Austin, Texas 78712 Institute for Cellular and Molecular Biology, University of Texas, Austin, Texas 78712

International Review of Cell and Molecular Biology, Volume 274 ISSN 1937-6448, DOI: 10.1016/S1937-6448(08)02002-9

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2009 Elsevier Inc. All rights reserved.

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progression is dictated by the neural network controlling the hypothalamus including the cells that synthesize and release gonadotropin-releasing hormone (GnRH) and their regulatory neurotransmitters. Glutamate and GABA are the primary excitatory and inhibitory neurotransmitters in the central nervous system, and as such contribute a great deal to modulating this axis throughout the lifetime via their actions on receptors in the hypothalamus, both directly on GnRH neurons as well as indirectly through other hypothalamic neural networks. Interactions among GnRH neurons, glutamate, and GABA, including the regulation of GnRH gene and protein expression, hormone release, and modulation by estrogen, are critical to age-appropriate changes in reproductive function. Here, we present evidence for the modulation of GnRH neurosecretory cells by the balance of glutamate and GABA in the hypothalamus, and the functional consequences of these interactions on reproductive physiology across the life cycle.

1. Introduction The development and maintenance of reproductive systems are necessary for the propagation of all vertebrate species. Each level of the reproductive hypothalamic-pituitary-gonadal (HPG) axis produces unique hormone(s) that regulate one another through feed-forward and feedback mechanisms. Not surprisingly, this system is remarkably complex and involves not only cross-talk among its components, but it also includes extrinsic inputs from other somatic and neural systems. This network needs to be considered in the context of the life cycle, as reproductive demands vary enormously through an organism’s life from birth to death. This review will focus on the neural (hypothalamic) control of reproduction across the life cycle, focusing on the female rat, a model for which most information is published. Because the hypothalamic control of reproduction is highly conserved among vertebrates, many of the conclusions drawn in the current article are likely to be in place in other species. However, when relevant we will note species differences. Because of the primary importance of the hypothalamus in driving reproduction, we will specifically discuss the roles of the hypothalamic neurons that produce the neuropeptide gonadotropin-releasing hormone (GnRH), and their regulation by two opposing neurotransmitter inputs: glutamate and GABA. Glutamate and GABA are the principal excitatory and inhibitory neurotransmitters, respectively, of the entire central nervous system including the hypothalamus (van den Pol, 2003), so it is not surprising that they play key regulatory roles in the control of GnRH neurons. Numerous other neuroactive substances are also critically involved in integrating the function of GnRH neurosecretion and we refer readers to other papers on this subject (de la Iglesia and Schwartz, 2006; Gore, 2002; Malyala et al., 2005), as it is far beyond the scope of the current review to provide a comprehensive understanding of

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the more than 50 neurotransmitters, neurotrophic factors, and other regulatory molecules that affect the GnRH system. Here, the interactions of GnRH neurons, GABA, and glutamate in the hypothalamus will be considered at molecular, anatomical, and physiological levels. In addition, the potential role of glutamate and GABA neurons in the mediation of feedback effects of steroid hormones on HPG function will be considered. There is no question that this hypothalamic network is an integral part of the overall circuitry that is responsible for the proper development, maintenance, and function of the reproductive axis in adulthood, and changes in this network may also contribute to its eventual decline through the life history.

2. The HPG Axis The capacity for vertebrates to reproduce is dependent upon a functional HPG axis. The hypothalamus encompasses a relatively small area of the brain at the ventral portion of the diencephalon, just below the thalamus. It contains neurons that release the decapeptide GnRH into the portal capillary system, where it is transported to the anterior pituitary gland, located just below the brain. Here, GnRH binds to its receptors on the gonadotropes and stimulates the release of the gonadotropins, luteinizing hormone (LH) and follicle-stimulating hormone (FSH). These are released systemically, and target their receptors on the gonads, where they bind and stimulate the release of steroid hormones (estrogens, progestins, and androgens in both females and males). These sex steroid hormones are transported through the general circulatory system, and as part of their activities, they feed back onto the pituitary and hypothalamus, inhibiting the release of GnRH and LH, completing a negative feedback circuit. Additionally, in females, just prior to ovulation, the estrogen-induced negative feedback changes to positive via a mechanism that is still not entirely understood but undoubtedly involves steroid-sensitive inputs to GnRH neurons. This causes stimulation of GnRH and LH, resulting in the preovulatory GnRH/LH surge that triggers ovulation in the females of species with spontaneous ovulation, including primates and rodents.

3. The Reproductive Life Cycle in Females 3.1. Development and the attainment of reproductive competence Throughout the lifespan, the reproductive axis undergoes a series of changes. During late prenatal/early postnatal life, the HPG axis is hyperactive in a sexually dimorphic manner, contributing to sexual differentiation of the

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brain (Gore, 2008; McCarthy, 2008). Later in postnatal life, this hyperactivity declines and the HPG axis enters a quiescent period during which GnRH release decreases, resulting in a decline of gonadotropin and steroid hormone levels. During the pubertal transition, the quiescent period ends, with basal GnRH levels, GnRH pulse frequency and pulse amplitude increasing, thereby activating the HPG axis, and resulting in the sexual maturation of the animal. This process includes the onset of folliculogenesis and ovulation in the female, spermatogenesis in the male, steroidogenesis in the gonads of both sexes, as well as the manifestation of reproductive behaviors. Although all three HPG levels become active during the pubertal period, the driving force is the hypothalamic GnRH neurons (Richter and Terasawa, 2001; Sisk and Foster, 2004). However, GnRH neurons have the ability to release the GnRH decapeptide long before puberty. Therefore, the attainment of reproductive competence involves not only molecular modifications of gene and protein expression of GnRH (Ebling and Cronin, 2000; Sisk and Foster, 2004), but also, a maturation of the hypothalamic neural circuitry that regulates the GnRH cells (Ebling and Cronin, 2000; Gore, 2002). Puberty involves changes in the excitatory and inhibitory neurotransmitters that regulate GnRH release, and of relevance to this article, is characterized by decreased GABAergic inhibition of GnRH neurons, and an enhancement of facilitatory effects of glutamatergic inputs to the GnRH system, among other neurotransmitters that modulate the timing of puberty (Clarkson and Herbison, 2006; Moguilevsky and Wuttke, 2001; Terasawa, 2005). In female rats, once puberty has occurred, the adult animal experiences regular estrous cycles of 4–5 days that are divided into four phases: diestrus I, diestrus II, proestrus, and estrus. During diestrus I and II, GnRH and LH release are low and estrogen (primarily estradiol) levels are at their lowest. As diestrus II transitions to proestrus, estrogen levels increase and concurrently switch from exerting a negative feedback onto the hypothalamus and pituitary, to positive feedback. This results in the GnRH/LH surge that triggers ovulation during estrus. The timing of reproductive behaviors is also coordinated by these cyclic hormonal fluctuations, with females becoming sexually receptive during this transition to estrus, the time that maximizes the likelihood of successful fertilization upon mating (Gore, 2008; McCarthy, 2008). Although this current article focuses on the rat model, a species that exhibits short estrous cycles (as do mice), other vertebrates, including nonhuman and human primates, undergo similar developmental processes, with the capacity to generate menstrual cycles first occurring during pubertal development. This latter process, while much more prolonged than the rodent’s estrous cycle, also shares similar traits of negative and positive feedback of preovulatory GnRH/LH release (Maffucci and Gore, 2005). Reproductive systems are subject to an array of biological clocks. A critical biological rhythm of reproduction is the pulsatile (circhoral)

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mode of release of GnRH, and subsequently LH. In rats, pulses of GnRH/ LH occur at intervals of roughly 30 min (Maffucci et al., 2008), and in primates, the rhythm is on the order of every 1–2 h (Terasawa, 1995, 2005). This mechanism is in place throughout the reproductive life of the organism and is necessary for proper reproductive function (Belchetz et al., 1978; Loucopoulos et al., 1984; Wildt et al., 1981). Notably, pulses of GnRH occur throughout reproductive cycles of females, and therefore are superimposed upon the longer cyclic changes in GnRH during estrous and menstrual cycles in rodents and primates, respectively. Even shorter biological rhythms of GnRH exist, as shown by electrophysiological recordings from GnRH neurons in explant cultures, with short (millisecond to second to minute) bursts of electrical activity, calcium currents, and fluxes in GnRH peptide. It has been proposed that the coordination of this activity results in the pulsatile GnRH pattern described above (Abe and Terasawa, 2005; Suter et al., 2000). The reproductive axis of mammals is also subject to circadian/ diurnal rhythms of 24 h (de la Iglesia and Schwartz, 2006). Additionally, at least some mammalian species exhibit seasonal rhythms of reproduction, and even more specifically, GnRH (Ebling and Cronin, 2000). Thus, in examining the development and maintenance of the female reproductive axis, all of these biological rhythms of reproduction must be considered. Here, we will focus on regulation of GnRH/LH pulsatile release and the preovulatory GnRH/LH surge.

3.2. Reproductive senescence During aging, reproductive systems undergo declines in function. While this is most apparent in the case of women, who undergo menopause at mid-life, animals also experience partial or total reproductive failure (Maffucci and Gore, 2005). Rats have proven to be an interesting model for reproductive aging because they experience neuroendocrine changes prior to ovarian failure. At middle age (about 1 year), the rat estrous cycle becomes irregular, resulting in cycles greater than 5 days in length. As the animal ages further, it eventually undergoes a transition to an acyclic state, resulting in persistent estrus, in which no ovulation occurs. This transition is accompanied by an attenuation and decline of the preovulatory or steroid-induced GnRH/LH surge (Wise, 1984), as well as a diminution of the GnRH/LH pulsatile pattern (Maffucci et al., 2008; Scarbrough and Wise, 1990). Additionally, changes to hypothalamic GnRH gene expression are seen (Bottner et al., 2007; Gore et al., 2000a, 2002; Rubin et al., 1997). Because these changes precede the loss of ovarian cyclicity and the capacity to ovulate, these data suggest that at least in rats, reproductive aging involves, and may even be causally linked to, a gradual failure of hypothalamic GnRH function. However, there may be substantial species differences in the mechanisms for reproductive senescence. Women undergo follicular loss at mid-life,

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making it very difficult to ascertain whether there are also neuroendocrine changes at this same time, not to mention the impossibility of measuring hypothalamic hormones in humans, as this is a highly invasive procedure. Such measurements have been made in the rhesus monkey, and Gore et al. (2004) showed that pulsatile GnRH release increases in perimenopausal rhesus monkeys, presumably due at least in part to a loss of negative feedback from estradiol. Consistent with this finding, serum LH, FSH, and gonadotropin free-a subunit (a proxy for GnRH) increased levels of their release during the perimenopause in women and female monkeys (Downs and Urbanski, 2006; Gill et al., 2002; Hall et al., 2000; Woller et al., 2002). However, later post-menopausal women do show a decrease in LH pulse amplitude and inter-pulse interval (Genazzani et al., 1997; Hall et al., 2000), suggesting immediate increases in gonadatropins due to the relief of estrogen stimulated negative feedback, but an eventual reversal of this elevation. Thus, while slight species differences may exist in hormonal profiles during this transition, likely due to the lack of ovarian degradation in the rat, overall trends in hormone release between aging rats and humans share some similarities. In aging rats, despite these changes in pulsatile and surge release of GnRH, and aberrations in GnRH gene expression, there are actually relatively few changes that are intrinsic to GnRH neurons. That is, GnRH cell numbers and distributions change very little with aging (Funabashi and Kimura, 1995; Hoffman and Finch, 1986; Miller and Gore, 2002; Miller et al., 1990; Rubin and King, 1994; Rubin et al., 1984; Witkin, 1986). GnRH neuronal ultrastructure does appear to undergo age-associated change (Romero et al., 1994), as does the association of GnRH cells with astrocytes (Cashion et al., 2003). Thus, we invoke the broader GnRH neural network—the GnRH neurons together with their direct and indirect inputs from neurons, glia, and steroid hormone feedback—as an instigator of reproductive aging. This hypothesis is supported by observations of agerelated changes in function of hypothalamic neurotransmitters and neurotrophic factors, and expression of their receptors and steroid hormone receptors. Some of these changes may occur directly upon GnRH neurons, others upon the more extended circuit of cells that regulates the GnRH dendrites, perikarya, and neuroterminals more broadly. The net effect of these alterations with aging is a loss of drive upon GnRH release (Yin and Gore, 2006).

3.3. The GnRH-glutamate-GABA connection: An introduction Glutamate and GABA are regulators of developmental-, adulthood- and age-related control of GnRH function. As the primary excitatory and inhibitory amino acid neurotransmitters, glutamate and GABA, respectively, are widely expressed in cells throughout the central nervous system.

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They each have several classes of receptors, which in turn, are made up of subunits whose expression varies by developmental age, sex, and region (Brann and Mahesh, 2005; Clarkson and Herbison, 2006; Gore, 2001; Henderson, 2007). Of relevance to reproductive neuroendocrinology, GnRH neurons themselves express glutamate and GABA receptors, and this coexpression varies across the life cycle (Adams et al., 1999; Bailey et al., 2006; Eyigor and Jennes, 1996, 1997; Gore et al., 1996; Jung et al., 1998; Miller and Gore, 2002; Sim et al., 2000). In addition, hypothalamic regions that mediate effects of steroid hormone negative and positive feedback are abundant in glutamatergic and GABAergic receptors. Here, we provide evidence for the control of GnRH function by glutamate and GABA, and how this regulation changes across the reproductive lifespan in females.

4. Glutamate and the HPG Axis Excitatory amino acid neurotransmitters, and their interactions with their receptors, are an extremely important mechanism of excitatory synaptic activity in the CNS. Of these, glutamate is the most abundant in the brain (Brann, 1995; Brann and Mahesh, 1994). Immunohistochemical and in situ hybridization studies examining glutamate receptor localization in the hypothalamus have shown their presence in a variety of hypothalamic nuclei, including medial preoptic nucleus (MPN), median preoptic nucleus, arcuate nucleus (ARC), dorsomedial, and ventromedial hypothalamic nuclei, supraoptic nucleus, suprachiasmatic nucleus (SCN), and paraventricular nucleus (Brann, 1995; Jennes et al., 2002). Additionally, immunohistochemical analysis of glutamatergic fibers in the hypothalamus has shown them to be in close proximity with GnRH perikarya in the preoptic area (POA) and GnRH axons in the median eminence (ME) (Eyigor and Jennes, 1996; Kawakami et al., 1998b).

4.1. Expression of glutamate receptors in the hypothalamus, and specifically on GnRH cells There are three classes of ionotropic receptors to which glutamate binds, named after agonist binding (Table 2.1). GnRH neurons express all of these: N-methyl-D-aspartate (NMDA) (Gore et al., 1996, 2000b, 2002; Gu et al., 1999; Ottem et al., 2002); kainate (Eyigor and Jennes, 2000), and DL-aamino-3-hydroxy-5-methyl-4-isoxazole proprionic acid (AMPA) (Bailey et al., 2006; Gu et al., 1999). Furthermore, these receptors are present not only on GnRH perikarya themselves, but also in a number of hypothalamic nuclei implicated in GnRH secretory control. These include anteroventral periventricular nucleus (AVPV), MPN, ARC, and ME, the latter being an

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Table 2.1 Ionotropic glutamate receptor classes and corresponding pharmacological agents Ligand Glutamate: endogenous ligand, binds to all glutamate receptors Receptor type

Receptor subunits

Agonist

Antagonist

NMDA NR1 receptor NR2a, 2b, 2c, 2d NR3a, 3b

NMDA (N-methylD-aspartate), NMA (N-methyl-D,Laspartate): specific agonists of NMDA receptors

AMPA GluR1–4 receptor

AMPA (DL-aamino-3-hydroxy5-methyl-4isoxazole proprionic acid): specific agonist of AMPA receptors Kainate: specific agonist of Kainate receptors

MK-801: noncompetitive NMDA receptor antagonist AP-5 or AP-7: competitive NMDA receptor antagonist Ifenprodil: Selective NR2b subunit antagonist NBQX: competitive antagonist to AMPA receptors DNQX: competitive antagonist to AMPA and kainate receptors DNQX: competitive antagonist to AMPA and kainate receptors

Kainate GluR5–7 KA1, receptor KA2

Other Glutaminase: Enzyme responsible for conversion of glutamine into glutamate Vesicular glutamate transporter (VGlut): Transporter of glutamate, the presence of which is thought to indicate a glutamatergic phenotype

area rich in GnRH nerve terminals (Brann and Mahesh, 2005; Gu et al., 1999; Yin et al., 2007). Activation of ionotropic receptors by glutamate has important effects on GnRH activity and resulting reproductive function, and this likely occurs via both these direct and indirect circuits. Conventionally, ionotropic glutamate receptors are designated as NMDA and non-NMDA (AMPA and kainate) glutamatergic subtypes. The NMDAR is believed to be a tetrameric receptor (Laube et al., 1998; Mano and Teichberg, 1998; Rosenmund et al., 1998), and is composed of multiple subunits (NR1, NR2a–d, NR3a–b; Table 2.1). In the hypothalamus, NR1,

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NR2a, and NR2b are abundant and are coexpressed on GnRH cell bodies (Eyigor et al., 2001; Gore et al., 2000b; Herman et al., 2000; Meeker et al., 1994; Miller and Gore, 2002). Non-NMDARs form tetrameric channels which, for AMPA-type receptors, may contain any of four subunits (GluR1–4), all of which are present in the hypothalamus and are coexpressed with GnRH neurons, although GluR1–2 appear to be the most abundantly expressed (Bailey et al., 2006; Eyigor et al., 2001). Kainate receptors may contain any of five subunits (GluR5–7, KA1, KA2) and all of these have been identified in rat hypothalamus, with KA2 being the most common (Eyigor et al., 2001; Meeker et al., 1994). The KA2 subunit is also the only such subunit to be colocalized immunohistochemically with GnRH perikarya in female rats to date, as shown by in situ hybridization ((Eyigor and Jennes, 1996). In addition, Todman et al. (2005) assayed the mRNA contents of GnRH neurons of adult female mice with PCR amplification and oligonucleotide microarrays. Of the ionotropic receptors, they detected the presence of NR2b, NR2d, GluR1, and GluR3, as well as Grid 1 and Grid 2, receptors of a fourth and less well known ionotropic glutamate class (d) that are not known to form functional channels in Xenopus oocytes or mammalian cells (Lomeli et al., 1993; Yamazaki et al., 1992). It should be noted that the lack of detection of other receptor subtypes does not necessarily reflect the absence of the mRNA, so these latter findings probably represent a subset of the total receptor subunit populations. As a whole, these immunohistochemical, molecular, and physiological assays indicate that GnRH perikarya have the capacity to respond directly to glutamatergic stimulation through a variety of ionotropic and metabotropic glutamate receptors. The previous discussion has focused on the expression of glutamate receptors at the level of the POA, both directly and indirectly, on GnRH cells. However, the GnRH neuroterminals also must be considered as a site of glutamatergic regulation. Using electron microscopy, Kawakami et al. (1998a) demonstrated in adult female rats that GnRH terminals in the ME coexpress NR1 and KA2 immunoreactivity. Although the GnRH terminals were not found to express GluR1, GluR2/3, and GluR6/7 in that study, other, non-GnRH, terminals in the ME were found to coexpress these receptors. The GT 1–7 cell line, an immortalized GnRH cell line that was produced using targeted tumorigenesis technique (Liposits et al., 1991; Mellon et al., 1991) also provides evidence for the expression of glutamate receptor subunits on GnRH neurons. As GnRH cells are often difficult to isolate and maintain in culture, these cells offer a viable alternative to studying GnRH neuronal function, and direct effects of glutamatergic influence on the action of these neurons. However, there is some disagreement in the field about whether and which glutamate receptors are expressed on GT1 cells. For example, the expression of the NMDA receptor subunits NR1 and

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NR2b–2d mRNA were reported (El-Etr et al., 2006; Mahesh et al., 1999; Urbanski et al., 1994). However, Mahesh et al. (1999) did not show the presence of GluR1, GluR4, and GluR6 mRNA, nor were NR1 or GluR1 protein present as determined by western blot analysis. Binding studies using AMPA, kainate, and NMDA specific antagonists suggested the absence or minimal effect of glutamate ionotropic receptors in this cell line (Mahesh et al., 1999). However, recent preliminary work from our laboratory (Maffucci and Gore, unpublished) did verify the presence of the NR1 and NR2a–2d subunit mRNA using real-time PCR, and both NR1 and NR2a (but not NR2b–d) protein using western Blot analysis. Furthermore, Garyfallou et al. (2006) found GluR2 and GluR4, as well as KA2 in the GT1–7 cell line using gene microarrays and RT-PCR. The variability of these data are likely due to differences in individual cell lines, emphasizing the importance of characterization of cell lines prior to experimental use. At the very least the results suggest the potential for GT1 cells to express glutamatergic receptors under certain culturing conditions. GT1 cells may also respond to glutamate agonists/antagonists. Incubation of these cells with AMPA, kainate, NMDA, or glutamate yielded increased GnRH release and an increase in intracellular calcium concentration (El-Etr et al., 2006; Spergel et al., 1994, 1995). Addition of the NMDAR antagonists MK-801 or AP-5 inhibited the NMDA-induced increase in GnRH release (Mahachoklertwattana et al., 1994).

4.2. Structure-function properties of glutamate receptors The structural properties of the ionotropic receptors determine their ability to bind and respond to glutamate. Specifically, the subunit composition of the receptor determines the kinetic properties of the channel, and thus understanding the stoichiometry of the receptor population, as well as its distribution in various nuclei in the hypothalamus, appears to be fundamental to understanding its influence on the reproductive axis. Of relevance to this article, the stoichiometry of glutamate receptors undergoes substantial developmental changes, having potential implications for glutamate’s target cells, including GnRH neurons. Moreover, the steroid hormone environment modulates the properties of glutamate receptors, making those cells that express the receptors potential targets of integration of steroid hormone influences on HPG function.

4.3. The glutamate-GnRH connection across the life cycle 4.3.1. Early development The role of the NMDAR on reproduction changes throughout the female’s lifespan. Some of these developmental differences may be attributable to either the absence of expression of the NMDAR in hypothalamic tissues,

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including on GnRH neurons, and/or the lack of a key receptor subunit that is required for the functionality of the receptor. This certainly appears to be the case during the infantile life stage. Our laboratory has quantified both gene and protein expression of NMDAR subunits in the preoptic areaanterior hypothalamus (POA-AH), a sexually dimorphic region that is important for reproductive physiology and sexual behaviors in male and female rats. Using rats across a range of early developmental ages, spanning late fetal development (day 18 of gestation) through early postnatal life (15 days of age), we reported age-, sex-, and subunit-specific differences in mRNAs of the NR1, NR2a, and NR2b subunits of the NMDAR in the POA-AH (Adams et al., 1999). Specifically, NR1 mRNA levels increased progressively during this developmental period, about 10-fold in males and 5-fold in females. NR2a mRNA was virtually undetectable in late embryonic life and remained extremely low in females through postnatal day (P) 15. In males, NR2a mRNA was first detectable on the day of birth and underwent a large increase from P10 to P15. Finally, NR2b mRNA underwent an approximate doubling in levels from P0 to P5 in both males and females. Taken together, these results suggest that the ability of cells to synthesize NMDAR subunits changes dramatically in the POA during early life development, a finding that has implications for differential abilities of NMDARs to respond to glutamate (Adams et al., 1999; Chen et al., 1999; Erreger et al., 2005; Vicini et al., 1998). We also evaluated whether GnRH neurons coexpress the NR1 subunit during early postnatal development and did not detect any coexpression at these ages. Finally, when the NMDA receptor agonist NMA, or antagonist MK-801 was administered to these developing rats, little effect on GnRH mRNA or primary transcript RNA levels was detected (Table 2.2; Adams et al., 1999). Thus, during the perinatal life period, the ability of glutamate to affect changes on the GnRH system is not due to direct actions via NMDARs on GnRH neurons, but rather, is either due to indirect inputs from other NMDAR expressing cells, or to non-NMDA glutamate receptors. 4.3.2. Puberty To our knowledge, the earliest studies demonstrating an involvement of glutamatergic NMDARs on HPG function came from reports determining consequences of NMDAR agonists on the timing of puberty in male rhesus monkeys and female rats. In those experiments, prepubertal animals were treated with an NMDAR agonist, and the results showed an advancement in the timing of pubertal landmarks (Gay and Plant, 1987, 1988; Urbanski and Ojeda, 1987). Since then and as elaborated upon below, studies have investigated pubertal changes in responsiveness of GnRH neurons to glutamatergic agonists/antagonists, through measurements of serum LH, central GnRH, and/or GnRH gene expression, in coordination with evaluation of external markers of puberty such as secondary sex characteristics.

Table 2.2 Effects of NMDA receptor agonists/antagonists on GnRH gene expression

Technique

a

Age, hormonal status

Steroid hormone treatment

Drug administration

Effect on GnRH gene expression

NMA (agonist) or MK-801 (antagonist) NMA twice daily

Little effects of NMA or MK-801 on GnRH mRNA or primary transcripta * GnRH mRNAb

RNase protection Intact female rat pups, assay postnatal days 0, 5, 10, 15

N/A

RNase protection Intact pubertal rats, assay beginning day 25 through the day of VO Northern blot Peripubertal (P28) OVX Hybridization rats

N/A

Estradiol/ progesterone

MK-801

+ in GnRH mRNA in rostral hypothalamusc

RNase Protection assay

a. Young intact (proestrus) b. Middle-aged intact

N/A

NMA

a. * GnRH mRNA in POA b. + GnRH mRNA in POAd

Quantitative RT-PCR

Proestrous female rats

N/A

MK-801

Attenuation of the increase in GnRH mRNA during the GnRH/LH surgee

In situ hybridization Real-time PCR

OVX adult rats

Estradiol

OVX young and middleaged rats

Estradiol

a. NMDA b. MK-801 Ifenprodil (NR2b antagonist)

a. * GnRH mRNA b. + GnRH mRNAf Little effect at either ageg

Adams et al. (1999). Gore et al. (1996). c Seong et al. (1993). d Gore et al. (2000b). e Suzuki et al. (1995). f Ottem et al. (2002). g Maffucci et al. (2008). Abbreviations: NMA, N-methyl-D,L-aspartate; OVX, ovariectomized; VO, vaginal opening; + decrease; * increase. b

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4.3.2.1. Glutamate synthesis and release in the hypothalamus increases during puberty The synthesis and release of glutamate in the POAhypothalamus changes during the life cycle, including puberty, and this may have functional outcomes on the reproductive axis. In vitro work by Bourguignon et al. (1995) suggested that an increase in glutaminase activity, the enzyme responsible for conversion of glutamine into glutamate (Table 2.1), occurred in conjunction with the observed increase in GnRH pulsatile release accompanying pubertal onset. Furthermore, glutamate levels in the MPN increased during the pubertal transition (Goroll et al., 1993). 4.3.2.2. Expression of glutamate receptors in the hypothalamus and on GnRH neurons increases during puberty NMDA and non-NMDAR expression in the hypothalamus undergo pubertal changes in mRNA and protein expression, and in the coexpression of their receptor subunits on GnRH neurons. For example, Gore et al. (1996) showed that NR1 mRNA levels in the POA-AH increased peripubertally, and using double-label immunohistochemistry, demonstrated that the coexpression of NR1 on GnRH neurons increased significantly during the time period when adult reproductive function was attained. Eyigor and Jennes (1997) used in situ hybridization to examine the coexpression of KA2 and NR2a with GnRH neurons in the rostral hypothalamus (medial septum-diagonal band complex and POA). Although they found no change in NR2a coexpression during the pubertal transition, KA2 was not only present in GnRH neurons, but showed a diurnal patterning throughout development, with higher expression levels in the morning from P20 to P40 switching to higher expression levels in the afternoon at P45–P50 (Eyigor and Jennes, 1997). The OVX, steroid-treated prepubertal rat has also been used as a model of glutamatergic regulation of GnRH function. Using in situ hybridization, expression patterns of GluR1–7 and NMDAR1 subunits in the AVPV, and changes in these patterns following estrogen or estrogen/progesterone treatment, were assessed (Gu et al., 1999). The results showed differential expression of these various receptor subunits in different structures within the AVPV itself, and little evidence for the presence of kainate receptors (GluR5–7). Furthermore, an estrogen dependent increase in the GluR1 subunit and suppression of NR1 (as compared to OVX controls) was reported (Gu et al., 1999) suggesting steroid regulation, a concept that will be discussed below (see Section 4.3.3.3). 4.3.2.3. Functional changes in the response of the HPG axis to glutamate during puberty Not only does glutamate receptor expression in the hypothalamus in general and on GnRH neurons in particular change during the progression of puberty, but the ability of glutamate to affect GnRH gene expression is facilitated during puberty. Gore et al. (1996) found that administration of the glutamate agonist NMA to intact female rats enhanced

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the naturally occurring increase in GnRH mRNA expression that accompanies the onset of puberty (Table 2.2). A further study examining blockade of NMDARs or non-NMDARs during the steroid-induced LH surge in peripubertal (P28) OVX, steroid replaced females showed that MK-801 (NMDAR antagonist), but not CNQX (AMPA-kainate antagonist), caused a decrease in the GnRH mRNA levels in the rostral hypothalamus (Seong et al., 1993). The ability of NMDAR agonists and antagonists to influence the reproductive axis does not become apparent until the pubertal time period. It appears that there is a critical period (P21 in rats) during which administration of an NMDAR agonist will induce precocious puberty in vivo and increase GnRH release in vitro (Moguilevsky et al., 1995; Smyth and Wilkinson, 1994). Prior to this age, NMDAR agonists/antagonists have relatively little effect on GnRH/LH release and the timing of puberty (c.f. Adams et al., 1999). Beginning at about P21, a time period when rats first begin to undergo activation of the HPG axis, the activation of NMDARs results in the induction of precocious puberty (Brann et al., 1993a; MacDonald and Wilkinson, 1992; Smyth and Wilkinson, 1994; Urbanski and Ojeda, 1987, 1990), while NMDAR antagonists delay pubertal onset (MacDonald and Wilkinson, 1990; Meijs-Roelofs et al., 1991; Urbanski and Ojeda, 1990; Veneroni et al., 1990). The functional role of non-NMDARs in this pubertal transition is less clear. Administration of kainate either systemically or directly (i.c.v.) into the third ventricle did not induce precocious puberty in immature female rats, nor did i.c.v. application of DNQX, an AMPA/kainate antagonist (Table 2.1), delay its onset (Brann et al., 1993a). Further investigation into why this difference exists between NMDARs and non-NMDARs suggest that these receptors may show sexually dimorphic mediation of puberty. Comparison of the activity of NMDA and MK-801 versus kainate and DNQX (AMPA/kainate antagonist) on gonadotropin secretion in both male and female prepubertal rats showed that, while the NMDAR drugs affected both sexes equally, kainate stimulated LH, and DNQX inhibited LH, in males only (Pinilla et al., 1998). Another study found slightly different effects on the gonadotropin response to kainate in males. While NMDA administration caused LH increases in vivo and GnRH in vitro in both males and females, kainate administration resulted in increased GnRH, but no effect on LH (Carbone et al., 1992, 1996). These differential findings in gonadotropin release are likely due to variations in methodology, including drug dose administered or time of day. However, overall, these results support a role for these receptors in the functioning of the reproductive axis during this important transition, and suggest a possible sexual dimorphism with kainate receptor action. Differential regulation of gonadotropin secretion by glutamate could be the result of differential distribution, and hence alterations in the stoichiometry, of the receptors

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within specific regions of the hypothalamus and their regulation of the GnRH neuronal populations, via direct or indirect mechanisms. More research will be required in order to better understand the physiological influence of glutamate and its various ionotropic receptors during this important developmental time period. It is important to point out that analyses of kainate and NMDAR binding in the hypothalamic area spanning the POA to MBH showed no change in the number of binding sites during the pubertal transition (Brann et al., 1993a). This finding suggests that the effects of these receptors are via increased sensitivity to glutamate or increased sensitivity of GnRH neurons to their activity, rather than to the ability of glutamate to bind to its receptors. Changes to receptor stoichiometries during this time period also need to be considered, as it is plausible that while receptor numbers remain the same, subunit distributions differ, changing the electrical properties and hence physiological responses of these channels. 4.3.3. Adulthood 4.3.3.1. Glutamatergic actions upon the GnRH/LH surge As mentioned previously, reproductive physiology in adult females involves several rhythms of GnRH/LH release. Of primary interest to reproductive physiologists studying species with spontaneous ovulatory cycles, such as rats and primates (both human and nonhuman), are the patterns of GnRH/LH pulsatile release and the GnRH/LH surge that precedes ovulation. Both of these events are influenced by the action of NMDARs and nonNMDARS. With respect to the activation of GnRH gene expression and release during the surge, NMDAR activation in proestrous young adult female rats (Gore et al., 2000b) led to rapidly increased GnRH gene expression in the hypothalamus (Table 2.2). The NMDAR antagonist MK-801 given to proestrous females not only blocked the LH surge, but also resulted in a less prominent increase in GnRH mRNA expression in the hypothalamus as compared to controls (Suzuki et al., 1995). However, in OVX, steroid hormone treated females, kainate injection (i.c.v.) administered on the morning of the expected steroid induced surge did not result in upregulation of GnRH mRNA levels, nor did it induce c-Fos expression in GnRH neurons 40 min following injection. However, it did result in an increase in LH levels 10 min following treatment (Eyigor and Jennes, 2000). Together, these data suggest that kainate is working indirectly to affect LH release during this time period. Furthermore, application of AP-7 (NMDA antagonist) or DNQX (AMPA/kainate antagonist) into the third ventricle of OVX, E2 replaced females blocked the steroid-induced LH surge (Lopez et al., 1990), as did administration of MK-801 to the proestrous female (Brann and Mahesh, 1991). Administration of NMDA or NMA on the morning of proestrus resulted in an increase in LH levels shortly after (Brann and Mahesh, 1992; Luderer et al., 1993). Thus, activation of glutamate

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receptors contributes to the onset of the preovulatory LH surge, with roles being played by both NMDA and non-NMDA receptors. 4.3.3.2. Glutamatergic effects on pulsatile GnRH/LH release Both NMDAR and non-NMDAR agonists and antagonists affect the pulsatile release of GnRH and LH in a similar manner to their effects on the GnRH/ LH surge. Agonists to both receptor types facilitate and antagonists diminish/block pulsatility. For example, pulsatile release of GnRH in vitro and LH release in vivo was diminished by administration of antagonists to NMDARs and non-NMDARs in both adult male and female rats (Arslan et al., 1988; Bourguignon et al., 1989b; Maffucci et al., 2008; Ping et al., 1994a, 1995). In vitro studies on explanted hypothalami have further implicated glutamate receptors as key regulatory factors of GnRH output. Application of AP-5 (NMDAR antagonist) and DNQX (non-NMDAR antagonist) to male hypothalamic retrochiasmatic explants resulted in an inhibition of GnRH output (Bourguignon et al., 1989a). Analysis of NMDA or kainate application to ME fragments from the OVX female in the absence of estrogen demonstrated similar increases of GnRH output (Kawakami et al., 1998a). Donoso et al. (1990) found that administration of the agonist kainate to ME fragments that had been explanted from male adult rats caused a 1.8-fold increase in GnRH output in vitro, while NMDA required a higher concentration (20 mM) to elicit a GnRH response. It should be noted that most of these latter studies focused on GnRH nerve terminals in the ME, a region where kainate receptors and NMDARs are expressed (Maragos et al., 1988; Unnerstall and Wamsley, 1983). In vivo experiments further highlight the role of these receptors in adult reproductive physiology. Kainate injected directly into the MPN of intact adult females on diestrus 1 resulted in increased GnRH content in OVLT and MBH, but gonadotropin output varied, with LH increasing, but not significantly, and FSH significantly decreasing (Gerendai et al., 1980). Repeated injections of kainate or NMA (i.v.) to intact (Abbud and Smith, 1991) or NMA (i.v.) to OVX (Pohl et al., 1989) female rats significantly increased LH release, but this elevation was not maintained with kainate. Again these findings support the potential for differential modulation of gonadotropin output based on the type of glutamate receptor activated. 4.3.3.3. Steroid modulation of glutamate’s effects on GnRH neurons The steroid hormone environment has considerable effects on glutamate receptor activation of GnRH neurons and has been studied at length in adult females. This is an important area of research because GnRH neurons do not express nuclear estrogen receptor (ER) a (Shivers et al., 1983) and although they express estrogen receptor b (ERb), this is not adequate to explain positive feedback effects of steroids upon the GnRH/LH surge (Wintermantel et al., 2006). Thus, effects of estradiol and progesterone on this event are mediated

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to a large extent indirectly by inputs to GnRH cells. Observations that hypothalamic inputs to GnRH neurons express ERa corroborate this idea. In the case of glutamatergic regulation, immunohistochemical analyses show that steroid hormone receptors (ERa, ERb, androgen receptor) are coexpressed with NMDARs and non-NMDARs in various nuclei of the hypothalamus that are important to regulation of GnRH release (Chakraborty et al., 2003a,b; Diano et al., 1997). Additionally, testosterone or estrogen replacement in gonadectomized adult males or females resulted in increases in GluR1 and GluR2/3 hypothalamic protein expression (Diano et al., 1997). By contrast, no effect of estradiol on NR1 immunoreactive cell numbers in the AVPV was found (Chakraborty et al., 2003a). It appears that the duration of estrogen treatment, as well as the time of day that the endpoints are examined, play a role in determining actions of glutamate receptors on GnRH output. In general, estradiol enhances NMDA-induced LH release (Arias et al., 1993; Gore, 2001; Maffucci et al., 2008). An examination of both GnRH and NMDAR subunit mRNA expression in response to different durations of estradiol treatment suggested little effect of duration in the POA-AH (Brann et al., 1993b; Gore et al., 2002; Ottem et al., 2002), but slight differences in the ME/medial basal hypothalamus (MBH) (Gore et al., 2002). Additionally, while NMDA administered in the morning (0700 h) increased GnRH mRNA levels in OVX, estrogen treated females, MK801 administered in the afternoon (1200 h) did not block the estrogen induced increase in GnRH mRNA levels that accompanies the estrogen-induced surge (Ottem et al., 2002). Effects of MK-801 administration in the morning were not examined. In focusing on AMPA receptors, Ping et al. (1997) found a differential effect of AMPA activation on GnRH release in vitro (MBH fragments from OVX, estradiol-primed or unprimed females) and LH in vivo (in OVX, estradiol-primed or unprimed females). The authors concluded that, while GnRH release increased regardless of the steroid hormone environment with AMPA administration and was reversed by administration of NBQX (AMPA specific antagonist), increases in LH were estrogen dependent. AMPA stimulated LH release in estradiol-primed females, but inhibited it in the absence of estradiol. Similar findings were seen when examining the effect of NMDA administration to OVX, steroid primed females (Arias et al., 1993). Thus, estradiol is a powerful regulatory factor in the modulation of this system by glutamate and its receptors. 4.3.3.4. Glutamate release in the hypothalamus-POA The ability of glutamate to activate GnRH release is clearly contingent upon its release at GnRH cell bodies, terminals, and dendrites. However, relatively little is known about this phenomenon in adult rats. Measurement of glutamate release in the POA shows that it is increased during the steroid hormoneinduced LH surge in adult female rats as measured by push–pull perfusion

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( Jarry et al., 1995; Ping et al., 1994b) and microdialysis sampling (NealPerry et al., 2005). These results are consistent with the proposed stimulatory role of glutamate during the GnRH/LH surge. 4.3.3.5. Conclusions on effects of glutamate on GnRH neurons of adults While these studies shed light on the role of these receptors, there are many questions yet to be answered. It appears that there may not only be differential effects of glutamate receptors on the reproductive axis, but that these effects may be mediated by the presence (or absence) of steroid hormones, and may be more or less robust depending upon the endpoint observed, including GnRH/LH pulsatility, the GnRH/LH surge, or other HPG axis responses at various time points during the cycle. The differential distribution patterns of these receptors throughout the hypothalamus suggest that specific nuclei may play different roles in regulating reproductive physiology in response to the presence (or absence) of glutamate. Thus, while we can conclusively state that these receptors play a role in the female reproductive physiology, the extent of that role, the individual contributions of each receptor, and their interplay, requires further research.

4.3.4. Reproductive senescence 4.3.4.1. Age-related changes in hypothalamic glutamate receptors Middle-aged rats experience a reproductive decline that results in decreased fertility and fecundity. Previous studies support that some of these changes are effected at the hypothalamic level and that excitatory amino acids are strongly implicated. Age-associated changes in expression of NMDAR and non-NMDARs in the hypothalamus have been shown that may underlie some of these functional differences. Gene expression of NR1, NR2a, and NR2b subunits were measured in the POA-AH and MBH of young, middle-aged, and aged intact animals of known cycling statuses (Gore et al., 2000b). NR2a and NR2b mRNA expression in the POA-AH were higher in cycling (both young and middle-aged) than acyclic (both middle-aged and aged) animals. In the MBH there was an overall increase in NR1, NR2a, and NR2b subunit mRNA expression with age. While this study showed no significant change in NR1 gene expression in POA-AH, by contrast, Zuo et al. (1996) found a decline in NR1 gene expression in both POA-AH and MBH regions in middle-aged versus young proestrus females using RT-PCR. The inconsistencies in findings may be due to the varied techniques used in each study, including tissue dissection, variation in times of sacrifice, mRNA extraction, and mRNA processing. However, both studies support the presence of important changes in NMDAR stoichiometry based on cycle status and age. Ovariectomized rats have also been used to examine age-related changes in gene and protein expression of NMDAR subunits. Using young, middle-aged, and old female rats that were OVX and given estradiol or

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vehicle treatment, Gore et al. (2002) showed significant effects of both age and hormone treatment on NMDAR subunit mRNA expression. Specifically, in POA, NR1 mRNA levels were significantly decreased by shortterm (2 day) but not longer-term (2 week) estradiol. NR2a mRNA levels were also down-regulated by estradiol, particularly in middle-aged and old rats. NR2b mRNA levels were highest in old rats and treatment with estradiol increased these levels in the old group. In the MBH, NR1 mRNA levels were decreased in estradiol compared to vehicle treatment; NR2a mRNA was not affected by age or estradiol; and NR2b expression was up-regulated by 2-week estradiol treatment in young rats, and decreased by this treatment in old rats (Gore et al., 2002). Therefore, these studies show the importance of regional specificity in the expression of these receptors, as results differed between the POA-AH and MBH. To further pinpoint the anatomical site at which estradiol may regulate NMDAR expression, the number of cells expressing the NR1 protein was quantified in the AVPV of OVX, estrogen or cholesterol treated aging female rats, a region chosen for its importance in the preovulatory GnRH/LH surge (Terasawa et al., 1980; Wiegand et al., 1980). Results of quantitative stereology showed no effect of estradiol, but significant ageassociated declines in NR1 immunoreactive cell numbers in the lateral AVPV (Chakraborty et al., 2003a). Recently, we have performed stereologic analyses of NR2b subunit expression in the AVPV of OVX rats at three ages (young, middle-aged, and old) and hormone treatments (Maffucci and Gore, unpublished; Fig. 2.1). Preliminary results suggest a decrease in NR2b-immunoreactive cell density in the AVPV from young to aged animals regardless of hormone treatment (vehicle–vehicle, estradiol–vehicle, or estradiol–progesterone). Furthermore, the volume fraction of the AVPV represented by immunofluorescent cells expressing colocalized NR1 and NR2b in this region increased with age, whereas the volume fraction of either NR1 or NR2b immunofluorescent independently showed no age associated changes (Fig. 2.1). Together, these data suggest that whereas the number of cells in the AVPV that express NR2b is decreasing with age, those cells that are remaining are more likely to coexpress NR1 with NR2b. The consequence of such changes would be altered channel properties of these receptors, and hence their effects on the reproductive system (Maffucci et al., unpublished). 4.3.4.2. Coexpression of glutamate receptor subunits on GnRH neurons and changes with age By examining NMDAR protein expression changes directly on GnRH neurons of the OVLT-POA hypothalamic region, our lab has demonstrated no age-related change in NR1 coexpression, but an increase in the NR2b subunit coexpression with GnRH from young to middle-aged intact animals (Miller and Gore, 2002). Thus, there may be a change in the stoichiometry of the NMDAR on GnRH cells with

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B

A 20 mM C

iiiV

200 mM

OT

Figure 2.1 (A) The anteroventral peroventricular nucleus is a region important to the integration of signals for proper female reproductive function. This region is located lateral to the third ventricle (iiiV) and rostral to the optic tract (OT), and is rich in cells immunopositive for the NMDAR subunits, including NR2b (shown here labeled by DAB-brown stain). A solid, black line highlights the border of the AVPV. The abundance of NR2b-immunoreactive cells can be seen in this representative middle-aged OVX rat treated with estradiol. (B) NR2b-immunopositive cells from another representative OVX rat (in this case, young vehicle treated) is shown at higher magnification (DAB-brown stain labels the NR2b cells). The Nissl counterstain is seen in blue. (C) Immunofluorescent labeling and confocal analysis of NR2b (green) and NR1 (red) in the AVPV of a representative rat (young, OVX plus estradiol plus progesterone) demonstrates the presence of colocalized (yellow) cells. The image size is 203 mm  203 mm.

aging, that may underlie some of the functional differences in the responsiveness of GnRH neurons to NMDAR agonists, as described below (see Section 4.3.4.3). Coexpression of NMDAR subunits on GnRH neurons during the steroid-induced LH surge, a time when the GnRH response to glutamate agonists in vitro is attenuated in middle-aged females. Zuo et al. (1996) showed differences between middle-aged and young rats. Specifically, AMPARs (GluR1, 3–4) on GnRH neurons underwent altered expression, as compared to young animals, on proestrus (Bailey et al., 2006). Specifically, GluR1/ GnRH coexpression was lower in middle-aged females than young at 1200 h and failed to decline at the end of the proestrus day (2000 h) in the young but not the middle-aged rats. GluR3 coexpression showed a diurnal pattern that was age specific. Young animals showed higher expression on the proestrus morning whereas middle-aged animals had a higher number in the afternoon.

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Finally, GluR4 coexpression in young animals was elevated early in the day as compared to middle-aged females, but showed no difference on the afternoon of the surge. These changing expression patterns likely contribute to the ageassociated differences in hypothalamic activity, and as discussed for NMDARs above, probably represent altered stoichiometries of non-NMDA receptors in the aging hypothalamus. 4.3.4.3. Functional changes in responsiveness of GnRH neurons to glutamate with aging Pharmacological studies addressed some of the functional consequences in changes in NMDAR stoichiometry and/or expression of NMDARs in hypothalamus and on GnRH neurons. In vitro application of NMDA to anterior POA-MBH fragments from young and aged OVX females resulted in increases in GnRH peptide concentrations, with young females showing a greater increase than the aged group (Arias et al., 1996). In this same study, in vivo administration of NMDA to the POA resulted in a greater LH response of the young animals compared to aged females, and baseline LH levels prior to NMDA administration were higher in the young animals (Arias et al., 1996). Zuo et al. (1996) performed a similar study using ARC/ME fragments from intact, proestrus young and middle-aged females. No age-associated differences were seen in basal GnRH output, but application of NMDA caused increases in GnRH, with a significantly greater effect in young explants. Furthermore, young animals had significantly higher levels of LH than middle-aged. However, this study did not measure the effect of in vivo NMDA application on LH levels. In assessing GnRH gene expression in vivo, NMA administration to intact young and middle-aged female rats resulted in an increase in young, but a decrease in middle-aged animals (Gore et al., 2000b). Thus, while GnRH release appears to be stimulated by NMDAR agonists, there appears to be an age-associated disconnection between release and GnRH mRNA, as well as an attenuation of the NMDA effect. Finally, a recent functional study of the role of the NR2b subunit in the reproductive senescent transition showed that inhibition of NMDARs containing the NR2b subunit with an NR2b-selective antagonist, ifenprodil, led to increases in overall LH levels and parameters of LH pulsatility in young and middleaged OVX, E2 treated females (Maffucci et al., 2008). In that study, little effect was seen on GnRH gene expression, suggesting that the action of ifenprodil is probably on the release of GnRH, independently of de novo GnRH biosynthesis (Table 2.2). Together, these results once again highlight the importance of NMDAR stoichiometry, particularly the presence or absence of an NR2b subunit, and suggests this subunit may cause inhibition of the GnRH neuron, either through indirect actions, or possibly through direct actions on GnRH neurons, as the NR2b subunit historically is associated with slower activation and deactivation channel kinetics (Chen et al., 1999; Erreger et al., 2005; Vicini et al., 1998).

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4.3.4.4. Age-related changes in hypothalamic glutamate release Glutamate levels in the POA decline during aging, as shown in vitro using male MPN-MBH explants (Bonavera et al., 1998) and in vivo using intracerebral microdialysis in steroid-primed females (Neal-Perry et al., 2005). Thus, beyond apparent changes to glutamate receptors themselves, there are accompanying changes in glutamate availability throughout the life cycle and it appears that these levels are regulated, at least in part, by the steroid hormone environment (Carbone et al., 1995; Jarry et al., 1992).

5. Gamma-Aminobutyric Acid (GABA) and the HPG Axis GABA is the dominant inhibitory amino acid neurotransmitter in the brain, including the hypothalamus, where 50% of all synaptic contacts in the hypothalamus are estimated to be GABAergic (Decavel and Van den Pol, 1990). It is derived from the neurotransmitter glutamate via the enzyme glutamic acid decarboxylase (GAD; Table 2.3), which is found in two isoforms, GAD65 and GAD67, of which GAD67 is viewed as the key enzyme for GABA synthesis (Asada et al., 1997). This relatedness between glutamate and GABA, the fact that they are the dominant excitatory and inhibitory amino neurotransmitters in the nervous system, respectively, along with other data presented in the next section, strongly suggests a close interplay between the glutamatergic and GABAergic regulatory pathways. As the key inhibitory amino acid in the adult hypothalamus, GABA is the main effector of inhibitory synaptic activity in this region (Decavel and Van den Pol, 1990). However, it has a very dynamic developmental progression. In addition to its role as a neurotransmitter, two of the GABA receptors (GABAA and GABAB; Table 2.3) have a large influence on the development of the central nervous system (Owens and Kriegstein, 2002). Anatomical studies of the these receptors and their respective subunit compositions during this period support a developmental change, as well as a shift in their physiological responses to the neurotransmitter (Clarkson and Herbison, 2006; Moguilevsky and Wuttke, 2001). Specific to the hypothalamus, studies support GABA, particularly through its action on the GABAAR, as affecting the establishment of sexually differentiated nuclei (McCarthy et al., 2002). It is important to note that the expression of the GABAC receptor appears to be primarily in the retina, superior colliculus, and cerebellum (Boue-Grabot et al., 1998) and, although this receptor has been detected in the pituitary gland (Boue-Grabot et al., 1995, 2000), it is not implicated in the establishment and maintenance of hypothalamic function. Taken together, these data suggest that, much like the glutamate receptors, the GABAergic system is undergoing developmental changes that have important repercussions on various neural systems, including the reproductive axis.

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Table 2.3 Ionotropic GABA receptor classes and corresponding pharmacological agents Ligand GABA (gamma-aminobutyric acid): endogenous ligand, binds to all GABA receptors Receptor type

Receptor subunits

GABAA a1–a6 receptor b1–b3 g1–g3 d, e, p, y gb1a GABAB receptor

Agonist

Antagonist

Muscimol; 4,5,6,7Tetrahydroisoxazolo [5,4-c]pyridin-3-ol hydrochloride (THIP): GABAA receptor agonists Baclofen: GABAB receptor agonist receptors

Bicuculline: competitive antagonist to GABAA receptors Phaclofen: antagonist to GABAB

gb1b Other Aminooxyacetic acid (AOAA): GABA transaminase inhibitor, inhibits GABA breakdown Glutamic acid decarboxylase (GAD): Enzyme responsible for conversion of glutamate into GABA GABA-transaminase (GVG): Enzyme that inhibits uptake and recycling of GABA GABA transporter (GAT): Transport protein, localized to neuronal and glial cells; provides rapid removal of GABA from the synaptic cleft Vesicular GABA transporter (VGAT): Transporter of GABA, the presence of which is thought to indicate a GABAergic Phenotype

5.1. Expression of GABA receptors in the hypothalamus and specifically on GnRH cells GABAA and GABAB receptors have been localized to many nuclei of the hypothalamus, both via mRNA (Whiting et al., 1997; Wisden and Seeburg, 1992) and protein (Davis et al., 2000; Fenelon and Herbison, 1995; Fritschy and Mohler, 1995) detection methods. Even more specifically, the a, b, and g subunits of the GABAA receptor have been colocalized to hypothalamic GnRH neurons in female rats ( Jung et al., 1998), and e subunits have been colocalized on GnRH cells in male rats (Moragues et al., 2003). Ultrastructural studies have provided evidence for GABAergic synapses on GnRH neurons in the MPN of the rat (Leranth et al., 1985). Thus, it appears that,

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like glutamate receptors, GABARs can influence the GnRH neuronal network directly as well as indirectly. The immortalized GnRH GT1–7 cells expresses GABA receptor subunits. The a and b, but not the g subunits, of the GABAAR have been detected (El-Etr et al., 1995; Favit et al., 1993; Garyfallou et al., 2006; Hales et al., 1992). Application of GABA to GT1 cells resulted in either a biphasic effect (an immediate stimulation of GnRH release followed by a suppression, suggested to be a result of two different GABARs working in concert) (Martinez de la Escalera et al., 1994) or solely a decrease in GnRH secretion (Favit et al., 1993; Martinez de la Escalera et al., 1994). As with experiments testing glutamatergic responses, these differential results are likely due to differences in growing and passaging of the cell lines used in either study. However, together, these in vivo and in vitro studies suggest a close link between GABA tone and GnRH function.

5.2. Structure-function properties of GABA receptors There are three types of GABA receptors (GABARs), two of which (GABAA and GABAC) are ionotropic and one of which (GABAB) is metabotropic. Both GABAA and GABAB appear to regulate the GnRH system. However, because the majority of research in rats has focused on the GABAAR, we will focus our discussion predominantly on the ionotropic receptor GABAA and mention other receptor types only when relevant. The GABAAR forms a pentameric structure for which 16 subunits have been identified (a1–a6, b1–b3, g1–g3, d, e, p, y; Table 2.3). Of these, a, b, and g are the most common to receptor composition (Henderson, 2007). Using recombination techniques, it has been demonstrated that in order to activate a functional chloride channel, the GABAAR must contain at least one a and one b subunit (Rabow et al., 1995; Wisden and Seeburg, 1992). The composition of the GABAAR is heterogeneous, regionally specific, and changes with age. The stoichiometry of this receptor, as with glutamate receptors, is indicative of its channel properties and its specificity to various GABAAR modulators. Additionally, specific subunits can also be modulated directly by steroid hormones, such as estrogens, having strong implications for the role of GABAAR stoichiometry in the regulation of reproduction (McCarthy et al., 1997).

5.3. The GABA-GnRH connection across the life cycle 5.3.1. Early development A key component of hypothalamic development involves alterations in GABA levels, receptors and synapses. Not only do GABA subunit expression patterns in the hypothalamus change with development, but the nature of the effect of GABA undergoes a developmental switch from excitatory to

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inhibitory. Thus, in early development, studies suggest that GABA stimulates neural developmental processes, and later in development (e.g., puberty), it appears to play inhibitory roles including the mediation of negative feedback effects of steroids on reproductive physiology. It is also important to note that although pharmacological studies support an inhibitory effect of GABA on GnRH release during puberty, electrophysiological studies actually suggest that GABA may exert excitatory effects on GnRH electrical properties that are potentially mediated through actions of GABA through a more complex neural network (DeFazio et al., 2002; Yin et al., 2008; Moenter and DeFazio, 2005). These changes in GABA levels, tone, and physiology appear to be key players in the regulation of GnRH neural circuits. Using radiolabeled dansyl chloride, a reagent that identifies primary amino groups, and hence is able to detect GABA levels, Cutler and Dudzinski (1974) showed an increase in GABA content in the rostral hypothalamus beginning at week 1 and going through to adulthood (please note that in this study the sex of the animals examined were not identified). Davis et al. (1999) expanded on these findings using HPLC to examine individual hypothalamic nuclei and found that these increases are nucleus specific, with no change occurring in POA, but increases in VMN and ARC observed in both developing males and females from P1 to P20. GABA plays a role in the sexual differentiation of the hypothalamus. Margaret McCarthy’s laboratory has shown the presence of sex and age dependent variations in GAD mRNA expression and GABA neurotransmitter levels in specific hypothalamic nuclei during the developmental time period (E18–P10) when organizational sex differences develop (McCarthy et al., 1997, 2002). Additionally, the GABAAR shows very interesting changes in both structure and function through development in the hypothalamus. In situ hybridization studies examining the distribution of a, b, and g subunit mRNA in male and female rats throughout early development (P0, 1, 3, 5, 6, 7, 10, 14, 21, 35, and adult, defined as 200–250 g) suggest a switch in the expression of the a1 and a2 subunits, with a1 showing an increase, and a2 a decrease in many forebrain nuclei, including hypothalamus. Additionally, it appears that the b3 subunit remains ubiquitously expressed throughout development (Laurie et al., 1992; Zhang et al., 1991, 1992). While these studies identified the distribution of many other receptor subunits, the developmental profiling was limited to a1, a2, and b3. Thus, other GABAAR subunits may change expression patterns with development, but have yet to be examined in this context. Examination of GABAAR protein expression of the a (1 and 2) subunit in both males and females from P1 through adulthood found developmental variations in the levels of the GABAAR-a1 subunit and GABAAR-a2 subunit, dependent on age and the hypothalamic nucleus studied (Davis et al., 2000). In the ventromedial nucleus (VMN), the GABAAR-a1 subunit decreased while GABAAR-a2 increased, whereas the POA showed no

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change in either subunit, and the ARC showed an increase in both. A different study found that 3H-muscimol binding (a GABAAR agonist; Table 2.3) in P20 animals was significantly higher as compared to earlier time points in the POA, but not VMN (Davis and McCarthy, 2000). Clark et al. (1997) detected an increase in binding in the VMN between P7 and P14, with a decline occurring by P70. These are not in complete disagreement with the findings of Davis and McCarthy (2000) as the latter showed a slight, but nonsignificant increase between P10 and P20. Together, these suggest that there is an increase in GABAAR activity during this transitional period, and an alteration is receptor activity based on nucleus specific changes in receptor stoichiometry. Expression of GABA subunits has also been studied specifically within developing GnRH neurons. Temple and Wray (2005) examined embryonic day (E) 11.5 mouse nasal explants at various time points (culture day 3.5, 7, 14, and 28) to determine the expression patterns of the GABAAR subunits a1–4, a6, and b3 using PCR analysis, and to ascertain their changes as a function of age. Some significant changes in expression were seen, suggesting alterations in the receptor stoichiometry during development in vitro. At culture day 28, GABAAR-a2 expression in GnRH neurons increased 20–33% from previous ages, to 75%, and a6 decreased from 83–100% to 25% at day 28. a1 maintained low expression levels throughout the time course (20–38%), while a3 remained at high levels (50–100%). Expression of a4 and b3 were variable (0–50%) but showed no significant differences. Thus, even at the embryonic stage, these subunits are modifying expression levels with development. The excitatory nature of GABA in early development is thought to influence neuronal growth, suggesting that this excitatory mechanism is functional during a time of neuronal migration, neurite outgrowth, and differentiation of various neurons (van den Pol, 1997). To examine the role of GABA on GnRH development, mouse embryonic olfactory tissues, in which GnRH neurons are present, were cultured and incubated with bicuculline, a GABAAR antagonist (Fueshko et al., 1998). Measures of neuronal outgrowth and migration showed no effect on GnRH cell number or fiber length, but did show an increase in the distance of neuronal migration (measured by the migration of GnRH-immunopostive cells from their olfactory pits). In that same study, incubation with the GABAAR agonist, muscimol, resulted in an inhibition of this migration (Fueshko et al., 1998). A recent live imaging analysis of GnRH neuronal migration (from embryonic mouse tissue) showed that application of bicuculline resulted in an increase in the percentage of video frames showing motion, and a decrease in the percentage of frames showing turns (Bless et al., 2005). This, in total, indicates that bicuculline application caused more forward motion, and less lateral motion. Together these two studies imply that the GABAAR plays a role in the facilitation of GnRH embryonic development and migration.

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5.3.2. Puberty There has been extensive research on the potential role of GABA as a mediator of puberty, with the long-held hypothesis that a removal of GABAergic input to GnRH neurons is necessary to enable the increase in pulsatile GnRH release that drives pubertal development. Evidence for this hypothesis has been provided by a number of different techniques, although there has been some disagreement in the literature as discussed below. It is noteworthy that push–pull perfusion measuring GABA levels in the POA of prepubertal (26 and 32 days) and post-pubertal (P45) female rats demonstrated a pubertal decrease in GABA (Goroll et al., 1993), consistent with a pubertal diminution of GABAergic tone on GnRH neurons. Examination of GABAAR gene and protein levels in GnRH neurons during pubertal development supports this trend of age-associated changes in GABA subunit expression. Jung et al. (1998) determined that the GABAAR subunits a1 and 2, b2/3, and g2 mRNA were all expressed within the GnRH neuron in female prepubertal (28 day) rostral hypothalamic regions (including medial septum, diagonal band of Broca, OVLT, POA, and rostral regions of the AH), whereas g1 mRNA was absent. In addition, Sim et al. (2000) compared juvenile (P15–20) and adult (P50–70) mRNA expression of GABA subunits in GnRH neurons of female mice and showed that juvenile GnRH neurons express a much more heterogenous population of GABAARs. a5, b1, and g2 were the most frequently coexpressed subunits, and excepting a4 and g1, all subunits that were probed (a1–5, b1–3, and g1–3) were detected. In general, these were detected at low frequencies in the juvenile hypothalamus, suggesting a very heterogenous population of receptors, whereas the adult hypothalamus showed much less variation. a1, a3, 5, b1, b3, and g2 were detected at high frequencies and specific combinations of these subunits were much more common. Additionally, more GnRH neurons coexpressed GABAAR subunits in the adult (Sim et al., 2000). It is noteworthy that GABABR subunits also change protein expression patterns in the hypothalamus during development. Western blot analysis of the GABABR-1a and 1b splice variants in female hypothalamic tissue at P1, P4, P12, P20, P28, and P38 showed an overall decline in both variants with age (Bianchi et al., 2005). Together, these findings support an anatomical change in the GABAergic system through development, both in the hypothalamus in general and on GnRH neurons in particular. To further understand how these anatomical changes may be represented by physiological output, electrophysiological studies examining the response of hypothalamic neurons to GABA stimulation or inhibition have been essential. Recordings taken from embryonic day 15 (E15) neurons in culture show that application of GABA induces an excitatory, rather than the expected inhibitory response in early neuronal cultures. This response gradually shifts to inhibition as the cultures age (Chen et al., 1995, 1996).

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Such a finding is consistent with results of Han et al. (2002). Here, GnRH neurons recorded from slice preparations of female immature (P10–17), peripubertertal (P25–30), and adult (P36–55) female mice showed differential responses to GABA application. Specifically, immature neurons showed a depolarizing response to GABA, peripubertal neurons exhibited a variable response of depolarization, hyperpolarization, or no response, and adult neurons were consistently hyperpolarized (Han et al., 2002). These findings suggest a developmental switch in the GABAergic response, in which initially, this system is stimulatory and then becomes inhibitory peripubertally. However, a study using similar recording techniques reported that GABA application actually depolarized GnRH neurons in adult (>42 days) female mice (DeFazio et al., 2002). A reconciliation of these results has not been made to date but differences may be attributable to direct versus indirect actions of GABA on GnRH cells (DeFazio et al., 2002; Moenter and DeFazio, 2005; Yin et al., 2008). These studies will be expanded upon below (Section 4.3.3.5). The translation of these electrophysiological data to the whole organism is crucial to understanding the importance of the GABA regulation of GnRH function. As mentioned previously, GnRH is released in a pulsatile manner, and this mechanism is functional in early postnatal life. However, in prepubertal animals, GnRH release enters a period of relative quiescence, during which the amplitude of pulses declines substantially, and then increase during the pubertal transition (Dohler and Wuttke, 1975; Hompes et al., 1982; Reiter and Grumbach, 1982; Roth et al., 1997). It has been suggested that the establishment of this quiescent period early in development is likely due, at least in part, to an increase in inhibitory GABAergic activity, and that the reversal of this process in the peripubertal period enables an increase in excitatory input to GnRH neurons. The period of the pubertal transition is one in which pharmacological properties of the GABAR undergo interesting changes. Activation of the GABA system using muscimol (a GABAAR agonist), baclofen (a GABABR agonist), or aminooxyacetic acid (AOAA: a GABA-transaminase inhibitor; see Table 2.3) peripubertally (P23–P30) resulted in an inhibition of GnRH release in vitro, LH release in vivo and the onset of puberty (Feleder et al., 1999). It appears likely that there is also a shift sometime between P20 and P30 whereby the GABAR, and more specifically the GABAAR, changes from an excitatory to an inhibitory action on the HPG axis (Brann et al., 1992; Moguilevsky et al., 1991; Szwarcfarb et al., 1994). As evidence, differential effects of muscimol and baclofen on GnRH in vitro and LH and FSH in vivo in prepubertal (P16) and peripubertal (P30) females have been demonstrated (Moguilevsky et al., 1991). While baclofen (the GABABR agonist) remained inhibitory on these HPG endpoints at both ages, muscimol (a GABAAR agonist) was stimulatory at P16, and inhibitory at P30. Brann et al. (1992), using estradiol-treated females, found that

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muscimol remained stimulatory on basal LH levels at P29 both in vivo and in vitro (the latter from hemipituitaries), but found no effect on GnRH release in vitro. These studies suggest that the pubertal period is a critical one for a shift in GABA receptor pharmacologic responses. Analysis of the muscimol (GABAAR) binding sites in female rats throughout development (P5–P45) showed that females have high levels of binding until the prepubertal timepoint (P15), at which point they experience a decline (Szwarcfarb et al., 1994). Additionally, administration of AOAA to females resulted in a stimulation of LH levels at P12, P16, and P18 with AOAA treatment, but a significant decrease at P30. In total, these data suggest that GABA binding to the GABAAR changes with development. However, further studies examining changes in GABABR binding are warranted to fully understand the physiology of these receptors, and their regulatory roles, during the pubertal period. 5.3.3. Adulthood The role of GABA in the adult female reproductive system has revealed both stimulatory and inhibitory GABAergic mechanisms that remain controversial to this day. An early pharmacological study in the OVX and OVX steroid hormone primed female rat showed an increase in LH release following GABA administration. In OVX females (steroid hormone primed were not tested), preinjection with bicuculline blocked the GABAergic rise in LH, suggesting an excitatory effect on this system (Vijayan and McCann, 1978). Using the GnRH GT1–7 cell line, Martinez de la Escalera et al. (1994) demonstrated a biphasic effect of GABA application, during which an immediate stimulation of GnRH release was observed, followed by a tonic inhibition. The application of specific GABAAR and GABABR agonists suggested that this stimulatory effect was GABAAR mediated, results that were replicated and expanded upon by Hales et al. (1994). An experiment by Virmani et al. (1990) used rat pituitary cells in culture, and applied GABA or muscimol either using a static incubation set-up, or a perifusion method with drug administration every 10 min. In the static setup, LH release increased in response to GABA or muscimol, whereas in the perifusion experiment, LH levels progressively decreased with application of both agents. Thus, pulsatile application of GABA and GABAergic drugs may prove more relevant in determining GABAergic function. In addition, these studies suggest direct effects of GABA on the pituitary level of the HPG axis, along with the considerable information showing hypothalamic actions of GABA on GnRH cells. 5.3.3.1. Pulsatile LH release The question of the role of GABA in the pulsatile release of LH has been extensively studied and results have been variable. This is likely due to (1) the models examined, including whether or not rats were OVXed, for how long, and if they received hormone

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replacement; (2) the dose of agonist or antagonist used; (3) the method of drug delivery; and (4) the brain region examined. To follow is evidence for the pulsatile release of GABA in the hypothalamus, as well as effects of pharmacological manipulations of GABARs on GnRH/LH pulsatile release. GABA release has been measured in the POA and MBH of adult female rats and shown to be pulsatile ( Jarry et al., 1988, 1992). However, the correlation of GABA pulses with LH (and presumably GnRH) pulses is debatable, as there is evidence for both the absence (Demling et al., 1985; Jarry et al., 1991) and presence ( Jarry et al., 1988) of such a correlation. This variability is likely due to the different techniques, or possibly different areas within the POA-AH, being probed. The POA-AH, the site of GnRH perikarya, has been a main focus of studies examining the effects of manipulations of GABA levels or receptors on GnRH/LH pulsatility. Administration of AOAA (Table 2.3) to OVX female rats resulted in a decline of mean LH levels, as well as the frequency of pulsatile events and the interpulse interval. Bicuculline (GABAAR antagonist) prevented this effect, and bicuculline also increased mean LH levels (Donoso and Banzan, 1984), suggesting a role for GABAARs in this mechanism. Complementary to that finding, muscimol (GABAAR agonist) treatment resulted in an inhibition of LH pulses (Akema et al., 1990). Furthermore, unilateral lesion of the POA resulted in a temporary (2 h) decline in mean LH levels, LH amplitude, and the number of pulses in OVX females, which recovered following a 48-h period ( Jarry et al., 1993). Application of GABA (i.c.v.) into the POA-AH provided the same outcome ( Jarry et al., 1993). Taken as a whole, these data suggest a role of GABA in the regulation of GnRH/LH pulsatility, but support the idea that GABA is not the only neurotransmitter regulating this event, and hence even in the presence of GABA, compensatory mechanisms may be in place to recover lost pulsatile function. Two studies examining OVX female rats found similar GABAergic effects using bilateral ablation techniques, but reported unexpected results when administering bicuculline (Herbison et al., 1991; Jarry et al., 1991). Bicuculline resulted in a decrease in mean LH levels (in both studies) as well as a decrease in pulse amplitude and frequency ( Jarry et al., 1991) or an overall absence of pulses (Herbison et al., 1991). This may indicate a disinhibition of the pulsatile system, specifically in reference to the GABAAR, whereby inhibition of this receptor causes stimulation of an inhibitory response. The interplay of this receptor with GABABRs likely adds another potential explanation for this mechanism. Moreover, we remind readers of the previously discussed studies showing some stimulatory effects of the GABAergic system in GnRH/LH release, which may come into play to explain the inhibitory effect of a GABAAR antagonist on pulsatile LH release. Clearly, more research on the precise mechanisms by which GABA affects the HPG axis is warranted.

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The role of the GABABR in LH pulsatility is not as well-studied as that of the GABAAR. To our knowledge, there is only one study examining this receptor subtype on pulsatile LH release. Administration of baclofen (GABABR agonist; Table 2.3) to OVX female rats led to a very complex response of the pulsatile properties of LH release, and the extent of these outcomes were dose dependent (Akema and Kimura, 1992). To summarize, the authors found a temporary ablation of LH pulses, followed by an increase in pulse amplitude and inter-pulse interval. Biphasic and triphasic fluctuations in mean LH release were also seen (Akema and Kimura, 1992). In total, this evidence suggests that both GABAARs and GABABRs are involved in a complex regulation of GnRH/LH pulsatility, and likely influence different endpoints of hormone release. It would be interesting to examine the interplay of these two receptors in the regulations of pulsatile output, as it is likely that these two are interacting, rather than acting as separate entities. This likely could explain the differential results observed with bicuculline administration, as well as the complex response to baclofen administration. Finally, researchers must strive further to examine smaller nuclei in the hypothalamus, as evidence suggests a finely defined neuroanatomy and neurochemistry in these regions that may underlie very discrete and different physiological responses. 5.3.3.2. The GnRH/LH surge The literature concerning the role of GABA in the GnRH/LH surge suggests this neurotransmitter’s strong involvement in this physiological process. Studies examining GAD67 mRNA levels as an indicator for GABA synthesis during the preovulatory (Grattan et al., 1996; Herbison et al., 1992) or steroid induced (Curran-Rauhut and Petersen, 2002) LH surge overall support region specific declines in GAD67 on the afternoon of the surge (summarized in Table 2.4). Although these studies differ slightly in their findings for specific hypothalamic nuclei, likely due to differences in methodology, they all demonstrate an active involvement of GABA in the rostral hypothalamus in the regulation of the GnRH/LH surge. Specifically, a diminution of GABA biosynthesis precedes the generation of the large increase in GnRH that occurs during the surge. A study by Leonhardt et al. (2000) contradicts these findings, suggesting that GAD67 mRNA in the intact female increases during the afternoon of proestrus. However, these findings are likely due to their use of the whole POA rather than a focus on specific nuclei within this region, as well as the use of competitive RT-PCR rather than in situ hybridization, as was used in the former studies (Table 2.4). Additionally, findings by Herbison et al. (1992) suggest that GABA synthesis in the MPN and diagonal band of Broca specifically, as measured by GAD67 levels, changes only during the proestrous surge, and not throughout the estrous cycle, although multiple timepoints throughout the cycle should be looked at to further develop this timeline.

Table 2.4 GAD67 mRNA expression shows regional specificity of expression in the hypothalamus throughout the GnRH/LH surge Animal model

Technique

Preoptic area

DBB/OVLT

MPN

ME/MBH

Adult, intact, proestrus Adult, intact, proestrus Adult, OVX rats given estradiol Adult, intact, Proestrus

RNase protection assay In situ hybridization histochemistry In situ hybridization histochemistry Competitive RT- PCR

Microdissections**,a

PM +,a

No changea

No changea

Microdissections**,b

No changeb

+ PMb

Not examinedb

Microdissections**,c

AM *; PM +,c

AM *, PM +,c

Not examinedc

PM *,d

Not examined*,d

Not examined*,d

Not examinedd

* Indicates that this study used a macrodissection of the preoptic area, in which this nucleus, as well as other hypothalamic nuclei, were included. ** Indicates that these studies used microdissections, examining single nuclei located in the preoptic area. a Grattan et al. (1996). b Herbison et al. (1992). c Curran-Rauhut and Petersen (2002). d Leonhardt et al. (2000). Abbreviations: DBB/OVLT, Diagonal band of Broca, organum vasculosum of the lamina terminalis; ME/MBH, median eminence/medial basal hypothalamus; MPN, medial preoptic nucleus; OVX, ovariectomized; + decrease; * increase.

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In further considering the estrous cycle, examination of GABA levels in the whole hypothalamus of intact female rats showed a significant decline on the proestrus day, as compared to estrus and diestrus (Loscher et al., 1992). Further, regionally specific changes in GABA levels were demonstrated using both push-pull perfusion in the POA-AH (Demling et al., 1985; Jarry et al., 1988) and microdialysis in the MPN (Tin Tin Win et al., 2004) of OVX-estradiol primed females. These studies demonstrated a decrease in GABA levels just prior to the LH surge, whereas levels in the ME/MBH did not decline ( Jarry et al., 1995). It appears that the observed decline in GABA in the MPN is specific to the proestrous afternoon, as very few changes were seen throughout the rest of the cycle (Mitsushima et al., 2002). This decline in GABA prior to the GnRH/LH surge presumably works to disinhibit GnRH neurons, thus allowing the GnRH release to increase. This mechanism is likely to involve actions on GnRH biosynthesis and not on neurosecretion per se, as changes appear to be limited to POA-AH and are not seen in the ME. Other enzymes involved in GABA metabolism appear to be involved in the LH surge. Administration of GABA-transaminase (GVG; Table 2.3), an enzyme that inhibits uptake and recycling of GABA, to intact, proestrus females resulted in an inhibition of the LH surge and the absence of ovulation (Donoso and Banzan, 1986). Application of AOAA, which inhibits GABA breakdown (Table 2.3) reduced the amplitude of the steroid-induced LH surge (Donoso and Banzan, 1984), presumably by maintaining relatively high levels of GABA. However, unilateral administration of bicuculline followed by electrochemical stimulation of the MPN caused a decline of LH levels and an inhibition of ovulation by a mechanism that at least in part involves opiate receptors (Taleisnik and Haymal, 1997). This suggests that GABA regulation of the GnRH/LH surge involves both direct and indirect actions on GnRH neurons. The respective roles of the GABAAR and the GABABR on the GnRH/ LH surge remain largely unknown. Whereas direct and continual infusion of GABA into the MPN (Herbison and Dyer, 1991) and injection into the third ventricle (Morello et al., 1989) of proestrous females resulted in an inhibition of the LH surge, administration of bicuculline prior to GABA recovered it, suggesting a role for the GABAAR in this action. Local injection of muscimol into the third ventricle (Akema et al., 1990; Akema and Kimura, 1993) or MPN (Seltzer and Donoso, 1992) also inhibited the estradiol-induced LH surge, and bicuculline administration reversed this effect. These studies all support a role of the GABAAR in the LH surge regulation. For the GABABR, bilateral infusion of baclofen (GABABR agonist) at either a low or high dose into the third ventricle of OVX, estradiol primed females resulted in a delayed (low dose) or inhibited (high dose) LH surge (Akema and Kimura, 1991). Follow-up studies from that same laboratory, in which baclofen was administered to OVX females and LH pulsatile

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release monitored, showed that baclofen interfered with GnRH/LH pulsatility, causing an immediate disruption in pulsatility (low dose) or a decrease in pulse frequency and increase in amplitude (high dose). The authors suggest that the GABABR-induced inhibition of the LH surge may be in part due to an interference with the normal pulsatile rhythmicity (Akema and Kimura, 1992). In an attempt to understand how the GABAA and GABAB receptors work as a unit to regulate the surge mechanism, Adler and Crowley (1986) administered muscimol or baclofen to intact, estradiolþprogesterone primed females, resulting in a decline in LH levels during the LH surge. This decline was not recovered by the simultaneous administration of bicuculline when administered with baclofen, but was slightly recovered in those animals receiving simultaneous muscimol (however, not to the level of control animals). Together, these studies suggest that both the GABAAR and GABABR affect the action of the surge, but still leave unanswered to what extent, if any, these receptors interact to do so. Future studies should focus on revealing the roles of these receptors in the surge, specifically examining their role in the timing of the surge, its amplitude, and other physiological components. The timing of the LH surge in rats is predictable and tightly regulated, and coordinated to the time of day as driven by the circadian pacemaker. Of relevance, GABA regulation of the GnRH circuit is dependent upon the time of day. Morello et al. (1992) found that administration of GABA into the third ventricle of proestrus females at 1100 h inhibited the onset of the LH surge, whereas administration at 1300 h had no effect. Further examination implicates the GABAAR system as a timekeeper for the surge. Bicuculline administration produced similar effects as GABA, in that earlier administration (between 0900 and 1100 h) had no effect on the preovulatory LH surge, but administration at 1000–1200 h caused a premature surge (Kimura and Jinnai, 1994) with higher amplitude (Kimura and Jinnai, 1994). Funabashi et al. (1997) examined coexpression of GnRH neurons with c-fos during the preovulatory LH surge and the effect of bicuculline administration thereupon. The bicuculline-associated advancement of the surge was replicated, and c-fos expression on GnRH neurons in rostral hypothalamic nuclei did not differ between those tissues collected in late afternoon from controls and early afternoon from bicuculline-treated animals, whereas those collected in late afternoon from controls had much lower expression levels. Thus, the GABAAR appears to act as a timekeeper for the GABAergic actions on the LH surge. To our knowledge, the role of the GABABR in this process is unknown. 5.3.3.3. GnRH gene expression Few studies have examined the role of GABA in GnRH gene regulation. Studies to date have suggested either no change, an increase, or a decrease in GnRH gene expression based on the

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pharmacology (GABAAR or GABABR) of the agonist or antagonist used in specific studies (Bergen et al., 1991; Cho and Kim, 1997; Kang et al., 1995; Leonhardt et al., 1995). These differences are likely dependent upon differing methodologies, absence or presence of steroid hormones, and dissection of the hypothalamic region. As summarized in Table 2.5, in OVX rats, GABA antagonists (both GABAA and GABAB) tend to increase, and agonists decrease, GnRH mRNA. When OVX rats are treated with estradiol, these GABA agonists tend to increase GnRH mRNA, an effect that is opposite of that in OVX rats that are not estrogen primed. Clearly, much more work is needed to clarify these interactions among GABA, estrogens, and GnRH neurons at the level of gene expression. 5.3.3.4. Steroid regulation of GABAergic systems As mentioned previously, the steroid hormone environment is critical to consider when examining the neurotransmitter regulation of the HPG axis. There is ample evidence to suggest steroid hormone regulation of GABA, beginning with coexpression of ERs with GABAergic neurons (as labeled by a GADspecific antibody; GAD-65 or GAD-67 not specified) in the POA-AH (Flugge et al., 1986). Furthermore, the GABAAR-a2 subunit is coexpressed with ER in the MPN and bed nucleus of the stria terminalis, and the expression of this subunit, along with GABAAR-b3, is regulated in these regions by estrogen (Herbison and Fenelon, 1995). Ovariectomy (OVX) in the adult female rat results in an increase in GnRH and LH basal levels due to the absence of estrogen-regulated negative feedback. In comparing GABA levels in the MPN of OVX female rats to intact, diestrous females, as measured by push-pull perfusion, levels were significantly lower in the OVX rats, consistent with the higher GnRH release (Ondo et al., 1982). Furthermore, bilateral infusion of GABA into the third ventricle of OVX females resulted in an acute increase in LH levels (within 30 min of this infusion); injection of bicuculline resulted in a slight and acute decrease. However, the animals treated with estrogen exhibited no such changes (McCann et al., 1984). Lamberts et al. (1983) replicated these findings in the OVX female, showing a decline in LH release following i.c.v. injection of muscimol, but no change in those treated with estradiol. These data, together, suggest that the presence of estrogen changes the nature of action of GABA on the HPG system. There is additional evidence that the presence of estrogen results in an increase in GABA levels in the POA-AH both in vitro (Herbison et al., 1989) and in vivo (Demling et al., 1985; Herbison et al., 1990; Mansky et al., 1982; Ondo et al., 1982), suggesting that GABA may play a role in the negative feedback effects of estrogens on GnRH cells. A more recent study examining the diagonal band of Broca (DBB), VMN, medial septum, MPN and dorsomedial nucleus (all nuclei within the rostral hypothalamus), OVX, estradiol treated females showed an increase in GABA levels in the DBB and

Table 2.5 Effects of GABA agonists/antagonists on GnRH gene expression

Technique

Age, Ovarian status

In situ Adult, OVX hybridization

Steroid hormone treatment

Drug administration

N/A

a. Bicuculline (GABAA antagonist) b. Tetrahydroisoxazolo[5,4-c] pyridin-3-ol hydrochloride c. Baclofen (GABAB agonist) (all into POA) a. Muscimol (GABAA agonist) b. Baclofen (GABAB agonist) (both into left lateral ventricle) Baclofen (GABAB agonist) (s.c. or into the third ventricle) a. Muscimol (GABAA agonist)

Competitive RT-PCR

Adult, OVX

N/A

Northern blot

Adult, OVX

Estradiol þ Progesterone

Northern blot

Adult, OVX

Estradiol þ vehicle or progesterone

b. Baclofen (GABAB agonist) (both into third ventricle) a

Effect on GnRH gene expression in POA

a. * GnRH mRNA b. No effect c. + GnRH mRNAa a. + GnRH mRNA b. No effectb * GnRH mRNAc a. * GnRH mRNA (low dose only) b. * GnRH mRNA (dose-dependent)d

Bergen et al. (1991). Leonhardt et al. (1995). Cho and Kim (1997). d Kang et al. (1995). Abbreviations: OVX, ovariectomy; THIP, Tetrahydroisoxazolo[5,4-c]pyridin-3-ol hydrochloride; POA, preoptic area; + decrease; * increase. b c

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VMN as compared to OVX controls (Luine et al., 1997). Furthermore, examination of intact, female rats suggests a relationship between estrogen and GABA levels throughout the estrous cycle, with GABA levels being highest in the medial POA during the morning of proestrus as compared to any other time in the cycle (Mitsushima et al., 2002). These data were expanded to suggest such correlation is regionally specific, with GABA levels varying on proestrus in the ventro- and dorso- medial hypothalamus only, and not in the MPN, diagonal band of Broca, lateral or medial septum, or AH (Frankfurt et al., 1984). Estrogen appears to selectively influence the enzymes involved in GABA synthesis, GAD65 and GAD67, and likely affects other regulatory proteins, such as GABA transporters (GAT; Table 2.3) responsible for removal of GABA from the synaptic cleft. Comparison of GAD67 mRNA level in the POA throughout the estrous cycle in females sacrificed at noon suggest no change in expression as measured by in situ hybridization (Herbison et al., 1992). However, as discussed previously, there are nucleus-specific changes in GAD67 gene expression during the preovulatory or steroid induced LH surge (Table 2.4). Further, findings from Curran-Rauhut and Petersen (2002) suggest an estrogen-mediated action during this period, as in situ hybridization studies examining the rostral POA/OVLT showed an increase in GAD67 levels on the morning of the estradiol-induced LH surge, and a decline in the afternoon, but no change in OVX females that were not treated with estradiol. GABA transporters (GAT) provide another source of possible estrogen regulation. Of these, GAT-1, expressed on neurons) and GAT-3, localized to both neuron and glial cells, are most abundant in the POA, and 75% of GATs in this region are estimated to be GAT-1 (Herbison et al., 1995). GAT-1 mRNA levels decline in a regionally-specific manner (specifically in the AVPV and MPN) and with OVX, whereby long-term (7-day) estrogen replacement resulted in significantly higher expression (in the MPN only). Furthermore, GABA uptake was enhanced, an unexpected finding that suggests the presence of estrogen is increasing transporter levels, the net result of which would be a decrease in extracellular GABA levels. In the caudal hypothalamus, Parducz et al. (1993) reported that the presence of estrogen resulted in ultrastructural changes to the GABAergic neurons of the ARC in the OVX, female rat. Specifically GABA-positive synaptic contacts on the membrane and axosomatic synapses declined with estrogen treatment. A similar study using the intact female across the estrous cycle, found a decline in GABA positive axosomatic synapses, and an increase in non-GABA labeled dendritic spine synapses during the transition from the proestrous morning to the proestrous afternoon and estrus (Csakvari et al., 2007). This suggests a diminution of GABAergic inhibition in this region and an increase in stimulatory synapses, thereby enabling female ovulation and reproductive behaviors during this time.

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Together, these studies show multiple levels at which estrogens and GABA may interact to influence the GnRH system. Apparently, all regulatory levels of GABA synthesis, release, and uptake are affected by estrogen, the net outcome of which is an apparent decrease in GABAergic tone to the GnRH cells prior to the preovulatory or steroid-induced GnRH/LH surge. We believe that this attenuation of GABAergic input then enables stimulatory inputs to GnRH neurons to come into play to stimulate the surge; a potential candidate for this stimulation is glutamate, among other neurotransmitters. We will discuss some of the possible interactions among GABA, glutamate and GnRH in Section 6. 5.3.3.5. Electrophysiological studies Electrophysiological studies focusing on the GABAergic drive on GnRH neurons have verified, without dispute, that GABAARs are located on GnRH neurons, that their activation results in functional physiological responses (Spergel et al., 1999), and that they are active at the synaptic level (Moenter and DeFazio, 2005; Sim et al., 2000; Sullivan and Moenter, 2003; Sullivan and Moenter, 2004, 2005). As mentioned previously, in early development (E15-peripubertal), electrophysiological recordings of non-GnRH, hypothalamic neurons (E15), and GnRH neurons in response to GABA application suggest an excitatory rather than inhibitory neuronal response (Chen et al., 1995, 1996; Han et al., 2002). However, in adult female mice, GABAergic activity was shown to be either inhibitory (Han et al., 2002) or excitatory (DeFazio et al., 2002; Moenter and DeFazio, 2005) on GnRH neurons, results that may differ due to several methodological differences: first, when considering the method of drug application, Han et al. (2002) applied bicuculline (GABAAR antagonist) as a bath. DeFazio et al. (2002) used a rapid application of muscimol (GABAAR agonist). The response of these cells to long-term versus shortterm application of drug likely varies, as receptors become activated and possibly desensitized to long-term application. Second, the method of GnRH identification also differed. In the former study (Han et al., 2002), GnRH neurons were identified using a GnRH-b-galactosidase transgenic whereas the latter study (DeFazio et al., 2002) used GnRH-GFP labeled animals. It is possible that these targeting techniques focused on two different populations of GnRH neurons that differed in their expression of GABA receptors in some way. Notably, a recent study (Yin et al., 2008) has also suggested excitatory effects of GABA on GnRH neurons of adult rats, consistent with work from Moenter’s laboratory. Research into other CNS regions (including cortex, SCN of the hypothalamus, and spinal neurons) support the idea that different populations of neurons may react to GABA differently, resulting in depolarization rather than the expected hyperpolarization (Gulledge and Stuart, 2003; Marty and Llano, 2005; Rohrbough and Spitzer, 1996; Wagner et al., 1997). Third, the former study (Han et al., 2002) did not block glutamatergic activity, whereas the latter

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study performed this blockade (DeFazio et al., 2002). As this review hopes to show, glutamate and GABA are two very important neurotransmitters that act both directly and indirectly to regulate this system and its inputs, as well as to regulate one another. The absence of glutamatergic signaling in one study versus its presence in another likely affects the final activity of the GnRH neuron. In another study, Moenter and DeFazio (2005), using hypothalamic tissue slices from female mice, showed that bicuculline, when applied in the absence of glutamate inhibition, caused a marked increase in the neuronal firing rate, whereas the presence of such inhibition resulted in a decreased firing rate, rather than the expected increase attributed to relief of inhibition. This work shows the importance of examining both the individual contribution of each neurotransmitter as well as their interactions. From these studies, it is important to recognize that there are likely different populations of GABAergic cells that respond differently to varying stimuli. The fact that GABARs contain many combinations of subunits and these are differentially modulated by steroid hormones (Herbison et al., 1995), that signals from other GnRH inputs (Roberts et al., 2008) integrate with GABAergic signals to affect a response, and that the expression of chloride transporters on GnRH neurons varies (Leupen et al., 2003, DeFazio, 2002) all suggest that the expected GABAergic result may vary depending upon the target tissue and the neural environment. 5.3.4. Reproductive senescence As discussed in Section 3.2, in many species including rats, aging is accompanied by a transition to a nonreproductive state. Part of this process involves changes to GnRH/LH pulsatile release, an attenuated and decreased GnRH/LH surge, and the change from a regular estrous cycle to a noncycling state. Despite abundant evidence for critical roles of GABA in the regulation of these mechanisms in young adults (described in the previous sections), it is surprising how few studies have examined GABA during the age-related transition to acyclicity. GABA turnover rates, using push-pull perfusion, were examined in intact young and aged female rats, and differences were observed in both the MPN and SCN, regions important to the modulation and timing of the GnRH/LH surge, ovulation, and circadian rhythmicity (Harney et al., 1996; Jarry et al., 1995). Acyclic, aged animals showed a decline in GABA turnover, as compared to young, in the MPN, but an increase in the SCN (in persistent estrous females only). Thus, changes to GABA turnover are likely affecting the regulatory mechanism of GnRH/LH release, such as pulsatile output, although further research is needed to resolve the mechanisms. Moreover, the net effect of decreased GABA turnover might be enhanced synaptic GABA. This possibility is consistent with a recent report by Neal-Perry et al. (2008) who showed by microdialysis that GABA

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release in the POA was increased in middle-aged compared to young ovariectomized, hormone-primed rats. In examining the GnRH/LH surge, Cashion et al. (2004) found that middle-aged animals had overall lower expression levels of GAD67 mRNA on the proestrus day, as compared to young rats, in the AVPV and organum vasculosum of the lamina terminalis (OVLT). Additionally, whereas in young animals GAD67 expression in these nuclei increased in the early morning of proestrus, followed by a decline at later time points, no such fluctuations were observed in their older counterparts in the AVPV only. Thus, the biological rhythms of GABA synthesis appear to be attenuated with age. A study by Grove-Strawser et al. (2007) supports these findings, showing a decline in GAD67 expression during the afternoon of proestrus in the OVLT/rostral hypothalamus of young female rats, but no such change in middle-aged females, and supporting a significantly higher expression level in middle-aged animals during the time period that LH is ascending. These data suggest a change in the synthesis of GABA such that the rhythmicity of its release, important to the timing of the preovulatory surge, is interrupted, thus providing one explanation for the alteration of this mechanism during the aging transition, contributing to an attenuation of the GnRH/LH surge. Consistently, the important study of Neal-Perry et al. (2008) confirms that pharmacological blockade of GABAARs with bicuculline in middle-aged rats mitigates the attenuation of the steroid-induced LH surge. These studies are just the beginning of numerous questions that researchers must examine to further understand the role of this inhibitory neurotransmitter in the reproductive senescent transition. Changes to GABA subunit populations, effects of agonists and antagonists, and their consequences on reproductive parameters (i.e., the preovulatory GnRH/ LH surge, GnRH pulsatile release, estrogenic regulation, prolongation of estrous cycles with age) need to be better understood to help shed light on the GABAergic mechanisms during this important life period.

6. Glutamate/GABA Interactions Thus, far we have examined the individual contributions of GABA and glutamate to the regulation of the reproductive system throughout the life span of female rats. These data show a very complex interaction of both GABA and glutamate in the regulation of the HPG axis, and suggest that these amino acid neurotransmitters set the inhibitory and excitatory tone for neurotransmission in the hypothalamus-POA (Fig. 2.2). As such, it is important not only to examine how they act individually to regulate this system, but also how they interact with one another in this regard. Although beyond the scope of this article, we point out that this glutamate-GABA

Prepubertal stage

Adult

Aged

Glutamate neurons (green)

GnRH neuron (yellow)

GABA neurons (tan)

Portal capillaries

Glutamate release in hypothalamus:

Low

High

GnRH co-expression of NMDARs:

Low

Increased c.f. prepuberty

Response to glutamate agon/antag:

Low

Change in NMDAR stoichiometry (especially NR2b) Excitatory, increased c.f. prepuberty Decreased c.f. adult

GABA release in hypothalamus:

High

Decreased c.f. prepuberty

GnRH co-expression of GABARs:

Co-expressed, with develop- Co-expressed, with changes mental changes in subunits in stoichiometry c.f. prepuberty

Response to GABA agon/antag:

Excitatory

GABA inhibits GnRH release, but causes mixed excitatory/ inhibitory effects on electrical activity

Decreased c.f. adult

Increased c.f. adult Unknown GABAA antagonist restores some aspects of the GnRH/LH surge

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balance in the POA provides a backdrop in which other hypothalamic neurotransmitters, neuropeptides, neurotrophic factors, and other neural factors then must act to exert their respective influences upon the GnRH neural circuit. Here, we provide a brief overview of the glutamate–GABA interactions across the reproductive life cycle. The glutamatergic and GABAergic systems are both in developmental transition during the early postnatal period (van den Pol et al., 1995). Indeed, during early development, these systems probably regulate one another in the hypothalamus to ensure a proper level of excitatory/inhibitory activity (Gao et al., 1998; van den Pol et al., 1998). van den Pol et al. (1995) used patch clamp recordings to determine the response of E15 rat hypothalamic neurons to GABA and glutamate agonists. These preparations showed that, while all measured neurons responded to GABA, only a fraction of these responded to the glutamate agonists (with no response recorded for NMDA application until E17). Later studies from this laboratory support this data and further suggest that the glutamate system develops more slowly than the GABAergic system, and that GABA may actually facilitate glutamatergic depolarization during this period (E15–E18) (Gao et al., 1998; van den Pol et al., 1998). This evidence suggests that during development, these systems have a very close interaction in which they regulate one another to insure a proper level of excitatory activity. The authors suggest that once established, the glutamatergic system becomes the main excitatory component and the GABAergic system switches to become the main inhibitory drive. From P15 to P30 there is a transition in hypothalamic GABAergic regulation, from a stimulatory to inhibitory action, and it appears likely that the GABAAR is the primary target receptor for this action. Furthermore, during this time the reproductive axis is undergoing a very important

Figure 2.2 Simplified model for interactions among GnRH, GABA, and glutamate neurons in the hypothalamus. For each of the prepubertal, adult, and aged life stages, a GnRH neuron is shown in yellow (terminals are shown containing large dense-core secretory vesicles of GnRH, in green); GABA neurons are colored tan and glutamate neurons are in green. At the prepubertal stage, GnRH neurons are depicted in relatively greater proximity to GABA neurons, with relatively little glutamatergic inputs. In adulthood, the balance between glutamate and GABA onto GnRH cells has shifted to a predominantly glutamatergic stimulatory influence. At this age, GABA likely provides a mix of inhibitory and excitory balance onto GnRH neurons, depending upon whether effects are direct or indirect. With senescence, this balance shifts in favor of inhibitory GABAergic inputs and a diminution of glutamatergic input to GnRH. Notably, at all ages, there are both direct effects of glutamate and GABA on GnRH cells as well as influences of glutamate and GABA cells on each other. Finally, while not shown, some cells may be both glutamatergic and GABAergic. Below each age is a brief summary of the relative effects of glutamate and GABA inputs on GnRH functions. Abbreviations: agon, agonist; antag, antagonist.

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change from quiescent to active, as the period from P25 to P30 represents the onset of puberty. It has been suggested that the GABAergic system interacts with the glutamatergic system during this transition to regulate the activity of the reproductive axis and influence these hypothalamic hormonal changes. Administration of AOAA, an inhibitor of GABA breakdown (Table 2.3) at P15 caused the expected increase in levels of GABA, and both glutamate levels in vitro and LH in vivo, followed, increasing as well. At P30, the AOAA-induced increase in GABA resulted in a decline in glutamate and LH levels, presumably due to the increase in GABA levels (Carbone et al., 2002). This suggests the possibility of GABAergic regulation of glutamate release, and a switch in that regulation from excitatory to inhibitory during the pubertal transition, causing a decrease in LH prepubertally, and an increase postpubertally. To further decipher the role of the GABAAR and GABABR in this, muscimol or baclofen was applied to hypothalamic explants of 30-day-old female rats. These treatments caused a decline in GnRH and glutamate release, suggesting that GABA, acting through both receptor types, may influence GnRH both independently as well as through an inhibition of excitatory glutamate release (Feleder et al., 1999). To determine the role of glutamate regulation of GABA activity during the prepubertal period, Scacchi et al. (1998) demonstrated that, while administration of the NMDAR antagonist MK-801 to prepubertal females did not significantly alter LH levels, treatment with the GABAergic agonists muscimol or baclofen increased or decreased LH levels, respectively, suggesting differential roles for the GABAAR (excitatory) and GABABR (inhibitory) during this time period. The addition of MK-801 to these GABAergic treatments resulted in an inhibition of the muscimol-induced LH increase, suggesting a robust regulation of the GABAARs by the NMDA system. Further, treatment with NMDA caused an increase in LH, which was inhibited by MK-801, but not bicuculline, implying that at this time period, the activity of the GABAARs is not regulating the NMDA system (GABABR activity was not measured). However, a study by Pinilla et al. (2002) found the opposite. Administration of either bicuculline or phaclofen (a GABABR antagonist) in P23 females inhibited NMDAstimulated LH levels. Although some of these results are not in agreement, possibly due to differences in methodology including different ages of experimental animals, mode of administration, or specific pharmacologic agent, the bottom line of these studies is that during pubertal development, there appears to be reciprocal interactions of glutamate and GABA, with downstream effects on the GnRH neurosecretory system. In the adult model, few studies have been undertaken to examine this interplay. Glutamate-stimulated release of GABA in various regions of the brain is well documented (de Mello et al., 1988; do Nascimento and de Mello, 1985; Harris and Miller, 1989; Moran et al., 1986; Pin and Bockaert, 1989). Examination of the POA specifically supported the presence of this

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mechanism, and further suggested that steroid hormone treatment, specifically estradiolþprogesterone, results in increased glutamate-induced release of GABA (Fleischmann et al., 1992) as measured from synaptosomes in culture. Using intact males, this same lab demonstrated that activation of GABAARs via muscimol (GABAA agonist) application resulted in both increased GABA and glutamate release from this same in vitro preparation (Fleischmann et al., 1995). An examination of a functional endpoint, such as the GnRH/LH surge, adds further insight to this observation. In OVX, estradiol treated rats given NMDA to facilitate the GnRH/LH surge, intracerebroventricular administration of muscimol (GABAA agonist), but not baclofen (GABAB agonist), resulted in an inhibition of the surge (Akema and Kimura, 1993). These results further suggest some interaction between GABAARs and NMDARs, but the nuances of this interaction are not known. Perhaps the most intriguing finding in this area is evidence for a dualphenotype GABA/glutamate neuronal population in the AVPV of the rat (Ottem et al., 2004). The AVPV is a nucleus of great importance to the execution of the GnRH/LH surge and ovulation. Ottem et al. (2004) used both in situ hybridization histochemistry and immunohistochemistry to show that the majority of neurons in the AVPV that express vesicular glutamate transporter (VGlut; Table 2.1), a marker of glutamate, also express vesicular GABA transporter (VGAT; Table 2.3), a marker of GABA, and vice-versa. Additionally, these dual-phenotype neurons strongly express ERa, they have contacts on GnRH neurons, and their glutamatergic-GABAergic phenotype is estradiol-sensitive. Specifically, during the estradiol-induced LH surge, the VGAT-containing vesicles decreased while VGlut-containing vesicles increased during the afternoon transition to the surge, suggesting an increase in glutamatergic, and decrease in GABAergic, activity during this event (Ottem et al., 2004). To our knowledge, this is the first model of its kind, and suggests an even more complex architecture for the regulation of the HPG reproductive axis by GABA, glutamate, and estradiol. Finally, to our knowledge, there is a single study that has interrogated the interplay between glutamate, GABA, and GnRH neurons during reproductive aging. Using ovariectomized young and middle-aged female rats, Neal-Perry et al. (2008) showed: (1) extracellular glutamate is decreased, and GABA is increased, in middle-aged rats with attenuated LH surges (measured by microdialysis); (2) pharmacological blockade of the GABAAR enhances both glutamate release and the LH surge; and (3) such treatment in concert with an inhibitor of synaptic glutamate reuptake further enhances glutamate levels and restores the LH surge. These results show a tight interaction of glutamate and GABA, along with GABAergic regulation of glutamate levels, and subsequent effects on the GnRH/LH surge (Fig. 2.2).

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7. Conclusion Understanding the reproductive axis, its development, maintenance, and senescence, has proven to be an important but difficult task due to the many regulatory components of this system. The complex interactions among the three levels of the HPG axis, as well as the neuromodulatory mechanisms upstream of this axis, creates an endocrine, neural, and neuroendocrine network that is only beginning to be fully appreciated. In considering the actions of each component part of this network, it is important to consider regional specificity of action, reproductive life cycle stage, and the steroid hormone environment, as many studies demonstrate differential mechanisms of modulation due to these variables. This review suggests that GABA, the main inhibitory neurotransmitter, and glutamate, the main stimulatory neurotransmitter, set a level of excitability in the hypothalamus that decreases or increases the likelihood that GnRH will be synthesized or released. On top of this tone, other neuropeptides and regulatory factors may modulate GnRH release in a manner appropriate to the animal’s reproductive status. For example, monoaminergic neurotransmitters such as serotonin, dopamine, epinephrine, norepinephrine; neuropeptides such as kisspeptin, neuropeptide Y, vasoactive intestinal protein, and many others; neurotrophic factors in the transforming-growth factor and insulin-like growth factor families, among others; and neuroactive gases such as nitric oxide; feed into this neurocircuitry. By further understanding the neural regulation of hypothalamic GnRH neurons, we will gain insights not only into normal reproductive functions, but better understand dysfunctions of puberty, infertility, and aging.

ACKNOWLEDGMENTS Grant support NIH AG16765 and AG028051 to ACG.

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Acquisition of Membrane Polarity in Epithelial Tube Formation: Patterns, Signaling Pathways, Molecular Mechanisms, and Disease Fernando Martı´n-Belmonte and Alejo E. Rodrı´guez-Fraticelli Contents 1. Introduction 2. Architecture of Polarized Cells in Epithelial Organs 2.1. Differentiated plasma membrane domains define epithelial architecture 2.2. Cell–cell junctions and polarity complexes 3. Epithelial Morphogenesis into Tubes 3.1. Model systems for studying tubulogenesis 3.2. Formation of epithelial tubes follows diverse patterns 4. De Novo Formation of Tubes: Conserved Pathways and Molecular Mechanisms 4.1. Initial cell–cell and cell–ECM interactions drive the formation of AJs and the orientation of the axis of polarity 4.2. Vesicle trafficking, membrane separation, and lumen coalescence in lumen formation 4.3. The role of PtdIns in the definition of membrane identity 5. Epithelial Polarity and Disease 5.1. Cell polarity and cancer 5.2. Trafficking disorders 5.3. Cytoskeletal and phosphoinositide related disorders 5.4. Cystic diseases of the kidney 6. Concluding Remarks Acknowledgments References

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Centro de Biologı´a Molecular Severo Ochoa, Consejo Superior de Investigaciones Cientı´ficas-UAM, Madrid 28049, Spain International Review of Cell and Molecular Biology, Volume 274 ISSN 1937-6448, DOI: 10.1016/S1937-6448(08)02003-0

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Abstract Epithelia coordinate the polarity of individual cells, in space and time, with surrounding cells and the extracellular matrix (ECM) to organize threedimensional structures that shape tissues and organs. One of the most important features of epithelial polarization is the asymmetric distribution of membrane surfaces with the apical surface facing a lumen or outside of the organism, and a basolateral surface facing other cells and ECM. This chapter discuss the processes required for the acquisition of the asymmetric distribution of membrane surfaces during morphogenesis, which include trafficking pathways, vesiclesorting machineries, formation of junctional and polarity complexes, and the establishment of signaling networks. In addition, different mechanisms and patterns are described for forming luminal spaces, and how alterations in cell polarity are associated with important human diseases such as cancer.

1. Introduction Many of our most important organs display a complex tissue organization made of a highly intricate network of branched tubes, which are required for the transport and distribution of liquids and nutrients throughout the body. Tubular organs develop following different strategies, which generate a remarkable structural and cellular diversity: different tube sizes, shapes, and connecting patterns (Hogan and Kolodziej, 2002; Lubarsky and Krasnow, 2003). These tubes are mainly composed of monolayers of highly polarized epithelial cells surrounding a central lumen. One of the most important features of epithelial cells is the existence of different plasma membrane domains, apical and basolateral, characterized by their distinct composition of lipids and proteins. Multiple cellular processes are required for establishing apical–basal polarity, including polarized vesicular transport, polarization of the cytoskeleton, and the proper establishment of cell adhesion and cell junction complexes. At specific stages of embryonic development and in some cancers, epithelial cells can undergo an epithelial-to-mesenchymal transition (EMT) that causes the loss of cell–cell adhesions and polarity markers, which results in the activation of a migratory phenotype. Therefore, the integrity of epithelial polarity plays an essential role in tumor progression and health (Debnath and Brugge, 2005; Yamada and Cukierman, 2007). Furthermore, many human diseases such as polycystic kidney disease, atherosclerotic heart disease, and faciogenital dysplasia derive from defects in epithelial tubular organization. A molecular understanding of tube morphogenesis could lead to new ways of diagnosing and treating these conditions. In this review paper, we focus on the acquisition of membrane polarity during epithelial tube formation. In particular, we will review different patterns that epithelial organs follow during morphogenesis, as well as the

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recent discoveries in the signaling pathways and the molecular mechanisms of these processes. Additionally, we will review how alterations in the process of acquisition of membrane polarity during morphogenesis are associated with important human diseases such as cancer.

2. Architecture of Polarized Cells in Epithelial Organs 2.1. Differentiated plasma membrane domains define epithelial architecture The epithelial plasma membrane is divided into three surfaces: the apical membrane, the free surface facing the lumen; the basal membrane, which is in contact with the tissues underneath; and the lateral surfaces that connect neighboring cells through a set of specialized cellular junctions that anchor cells to one another. These cellular junctions provide barrier function, and preserve the integrity and the different composition of the membranes by blocking the movement of proteins and outer-leaflet lipids (Mostov et al., 2003). The apical membrane is often covered by abundant microvilli, and appears to be the more specialized domain, since it contains most of the proteins required for organ-specific functions, such as terminal digestion and nutrient absorption or resorption. Generally, the apical plasma membrane is enriched in PtdIns-4,5-p2, sphingomyelin-containing glycolipids, cholesterol, and glycolipid-anchored proteins. By contrast, the basolateral membrane is enriched in PtdIns 3,4,5 p3 and carries most of the constitutive functions of the cells (e.g., cholesterol uptake, growth factor receptor, etc.). To maintain cell polarity and play their specific functions, epithelial cells have to ensure proper delivery of apical and basolateral cargos to their respective target location. The sorting of membrane proteins into distinct transport carriers occurs, either in the trans-Golgi network (TGN) during direct biosynthetic delivery, or in the common recycling endosomes (CRE) during indirect biosynthetic delivery or cargo-recycling to the plasma membrane (Folsch, 2008; Rodriguez-Boulan et al., 2005). Polarized protein delivery is regulated by sorting signals contained within the proteins themselves, which are recognized by specific sorting machineries. Some of the components of these specific sorting machineries have been identified in recent years, providing crucial information about the molecular mechanism that govern the protein traffic in epithelial tissues (Folsch, 2008; Mostov et al., 2003; Rodriguez-Boulan et al., 2005). Figure 3.1 summarizes the different trafficking routes and sorting mechanisms in epithelial cells that include: biosynthetic, endocytic, recycling, and transcytotic. Biosynthetic routes provide newly synthesized proteins to the apical and basolateral membranes. After the synthesis of the proteins, they are transported along

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Figure 3.1 Trafficking pathways in polarized epithelial cells. Biosynthetic pathways allow the transport of newly synthesized proteins to the apical and basolateral surfaces from the TGN. The biosynthetic pathway to the basolateral surface could be direct or indirect through the common recycling endosome (CRE). See Folsch (2008) for details. Apical and basolateral endocytosis allows the internalization of surface proteins to the early endosomes; the apical early endosome (AEE) or the basolateral early endosome (BEE). The proteins at the early endosomes can be: (A) recycled back to the surface using the recycling pathways at the apical, through the apical recycling endosome (ARE), and/or basolateral surfaces the CRE; (B) send to the opposite surface using the pathway for transcytosis from the basolateral to the apical surface; (C) or send to the late endosomes (LE) for degradation.

the secretory pathway (ER, Golgi, and TGN), and sorted into carriers to the different domains at either the TGN, or the endosomes. Proteins at the plasma membrane are rapidly endocytosed and delivered to the early endosomes where they follow the endocytic route or, after passing through the CRE, are recycled back to the cell surface (recycling), or transported across the cell to the opposite plasma membrane (transcytosis). The importance of these pathways varies with the type of epithelial cell, but they must be finely regulated in order to induce and maintain the steady-state polarity of the cells. The transport of proteins along these trafficking routes is regulated by sorting signals present in the proteins themselves and recognized by specific sorting machineries.

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2.1.1. Sorting signals and sorting machineries It was initially proposed that whereas the basolateral sorting was dependent on cytoplasmic signals, the apical sorting was a ‘‘default’’ pathway. This hypothesis originated in experiments in which the mutation of the basolateral signals (described below) induced the missorting of the proteins to the apical domain (Matter and Mellman, 1994). However, further work has proved the existence of apical sorting information that typically consists of ectodomain, membrane, or cytoplasmic signals. The first apical signal characterized was the glycosylphosphatidylinositol (GPI) anchors present in certain proteins (GPI-anchored proteins) (Lisanti et al., 1988). The evidence for these results came from a series of experiments in which the recombinant addition of the GPI anchor to secretory proteins resulted in the apical localization of the chimeric proteins (Brown et al., 1989; Lisanti et al., 1989). A second group of apical sorting signals includes N-linked or O-linked glycans, present in the exoplasmic region of many glycoproteins. A third group of apical sorting signals is encoded by the protein sequences themselves (Rodriguez-Boulan et al., 2005). One example is the transmembrane domain of the influenza virus protein hemaglutinin (HA), a prototypical apical targeted protein, which contains apical sorting information (Tall et al., 2003). Finally, it has been recently described that apical sorting information can also be encoded by cytoplasmic and exoplasmic protein domains present in the apical proteins (Marzolo et al., 2003; Takeda et al., 2003). A common requirement for apical sorting seems to be a clustering of the apical proteins into specific membrane domains, perhaps with the help of lectins that recognize N- or O-linked glycans, for direct delivery from the TGN to the apical membrane (Delacour et al., 2005; Fiedler and Simons, 1995) or, due to the ability of some apical directed proteins such as GPI-anchored proteins, to oligomerize during their passage through the Golgi complex (Paladino et al., 2004). Additionally, these apical clustering could be mediated by either lipid-raft domains or nonraft carriers. The lipidraft hypothesis (van Meer and Simons, 1988) postulates that apical targeted proteins are clustered and incorporated in transport vesicles due to their affinity for microdomains enriched in glycosphigolipids and cholesterol. Different proteins have been postulated to promote the clustering of lipidrafts, including the MAL/VIP17 (Alonso and Millan, 2001) and the caveolin family of proteins (Simons and Ikonen, 1997), which induce the formation of large amounts of cytoplasmic membrane vesicle structures when overexpressed in insect cells (Li et al., 1996; Puertollano et al., 1997). The association of the MAL/VIP17 family of proteins with lipid-rafts and their biological function seems to be related to its MARVEL domain, which is also present in physins, gyrins, and occludin families (Sanchez-Pulido et al., 2002). The function of the MARVEL domain could be related to cholesterol-rich membrane apposition events in a variety of cellular

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processes, such as biogenesis of vesicular transport carriers or tight junction regulation. The role of MAL/VIP17 and other members of the family in raftassociated vesicle transport has been extensively addressed in epithelial cells (Cheong et al., 1999; de Marco et al., 2002; Martin-Belmonte et al., 2000, 2001, 2003; Puertollano et al., 1999). However, the specific role for caveolin protein in raft-associated vesicle traffic still needs to be clarified (Manninen et al., 2005; Scheiffele et al., 1998). Some apically transported proteins, such as gp114, the neurotrophin receptor p75, and lactase-phorizin hydrolase (LPH), are included in nonraft vesicle carriers (Delacour et al., 2006). These nonraft vesicles, however, need also to be clustered for apical delivery. In fact, recent data has shown that Galectin-3 can form oligomers, and is involved in the clustering and apical delivery of nonraft associated glycoproteins to the apical surface and to membrane polarization of mouse enterocytes in vivo (Delacour et al., 2006, 2007, 2008). Interestingly, it appears that nonraft and raft dependent carriers require the presence of adaptor proteins at the TGN for apical delivery. The phosphatidylinositol(4)phosphate (PI(4)P) binding protein FAPP2 has been recently described to be required for apical delivery of the raft-associated protein HA and GPI-anchored proteins, and also for nonraft-associated vesicles (Vieira et al., 2005, 2006). FAPP2 seems to be needed for the transport of glycosylceramide from the cis-Golgi to the TGN, where it is incorporated into glycosphingolipids (D’Angelo et al., 2007). The authors speculate with the possibility that FAPP2 coordinates glycosphingolipid synthesis with apical transport. The basolateral sorting information, by contrast, is composed of sorting peptides included in the cytoplasmic tails of the basolateral targeted transmembrane proteins. Basolateral sorting signals, first described for the polymeric IgA receptor (pIgR) (Casanova et al., 1991; Mostov et al., 1986), are typically tyrosine-based (YXX) or leucine-based (mono- or di-leucine) peptide motifs (Rodriguez-Boulan et al., 2005). They are usually dominant over apical sorting signals, which explain the fact that when they are removed, certain basolateral proteins are missorted to the apical surface (Folsch, 2008). These peptidic sorting signals are recognized by cytosolic adaptor proteins, which in general form heterotetramers and interact with the vesicle coating protein clathrin, also required for basolateral distribution (Deborde et al., 2008). There are four types of complexes AP-1, AP-2, AP-3, and AP-4, and five types of adaptin proteins. Each complex consist of two large subunits (a, g, E, d, and b, 1
4), a medium subunit (m, 1–4), and a small subunit (s, 1–4). The tyrosine sorting signal (YXX) is recognized by the m subunit, whereas the di-L motif is recognized by the g/s1 or d/s3 subunits ( Janvier et al., 2003). The interaction of the adaptor proteins with the cargo and the clathrin coats induces the clustering of the basolateral proteins into clathrin-coated pits, which are subsequently budded into the cytoplasm for basolateral distribution. However, the current knowledge

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about the specificity of the adaptor proteins in basolateral trafficking is still very poor. Furthermore, the only AP complex that has been proven to be associated with basolateral sorting identified so far is the AP–1B complex. The AP–1B complex is composed of the m1b subunit, that determines exclusively the polarity of tyrosine-based signals, and it is only expressed in a number of polarized epithelial cells (Folsch et al., 1999; Gan et al., 2002). The current knowledge about the transcytosis signals is very limited, except for pIgR, which has been extensively studied for many years (Mostov et al., 2003). Recent findings suggest the possibility that the inactivation of basolateral sorting signals, when the proteins have already reached the basolateral surface could induce the activation of apical delivery signals, and thus, mediate the transcytosis of proteins that present both sorting information from the basolateral-to-the-apical surface (Anderson et al., 2005). Furthermore, this potential mechanism for transcytosis would be defined by the dominant hierarchy of basolateral signals over the apical signal described before, which would allow the initial delivery of the proteins to the basolateral domain. 2.1.2. Membrane fusion machinery The tethering (or docking) and fusion of transport vesicles with apical or basolateral domains are essential during exocytosis, and are needed for the acquisition of cell polarity and membrane identity (Wu et al., 2008). However, little is known about the mechanisms that control all these events during exocytosis. The exocyst protein complex function has been reported to be involved in the tethering, docking, and fusion of post-Golgi vesicles with the plasma membrane in polarized cells (Fig. 3.2). The most recent information from several model systems has demonstrated that the three families of small GTPases (Rab, Ral, and Rho) regulate the function of the exocyst complex, which is composed of eight subunits conserved from yeast to mammalian cells: Sec3, Sec5, Sec6, Sec8, Sec10, Sec15, Exo70, and Exo84 (Wu et al., 2008). The initial vesicle-docking event is regulated by Rab and Ral GTPases, perhaps by promoting exocyst assembly. There is evidence that the exocyst assembly is regulated by Ral and this function, like that of Rab GTPases, is first required for vesicle tethering rather than fusion (Moskalenko et al., 2002, 2003). Assembly is followed by local activation of the exocyst complex by active, GTP-bound, Cdc42 or TC10 GTPases. Exocyst activation results in the stimulation of a downstream fusion activity, probably by promoting assembly of active t-SNARE heterodimers. The presence of active t-SNARE proteins results in the fusion of the secretory vesicles at the site of exocyst activation. In epithelial cells, the exocyst is localized in the Golgi apparatus, the TGN, RE, and the junctional complex, and is proposed to promote the targeting and fusion of biosynthetic and endocytic recycling cargo carriers with the basolateral plasma membrane domain, possibly at sites near the tight junction (Folsch et al., 2003;

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Figure 3.2 Model for basolateral vesicle docking and fusion regulated by small GTPases and the exocyst. See Wu et al. (2008) for more information. The initial vesicle-docking or tethering event is regulated by Rab8 and RalA GTPases by promoting exocyst assembly. The association of the exocyst subunits with the vesicle or plasma membrane in this diagram is speculative. This is followed by local activation of the exocyst complex by GTP-bound Cdc42 Rho GTPase. Exocyst activation results in a stimulation of downstream fusion activity, probably by promoting assembly of active t-SNAREs heterodimers (SNAP23 and Syntaxin 4). The presence of active t-SNARE dimers with ts-VAMP, the v-SNARE, results in SNARE-mediated fusion of the vesicles with the basolateral membrane at the site of exocyst activation.

Grindstaff et al., 1998; Yeaman et al., 2001, 2004). More recent results have shown that the exocyst could function in several endocytic pathways as well, including basolateral recycling, apical recycling, and basolateralto-apical transcytosis. The latter was selectively dependent on interactions between the small GTPase Rab11a and Sec15A (Oztan et al., 2007). In the apical membrane, current information about the machinery for vesicle fusion is limited. The annexins family of proteins, which associate with the plasma membrane in a Ca2þ and negative phospholipids dependent manner (Rescher and Gerke, 2004), could be part of the machinery that mediates the fusion of apically derived vesicles with this plasma membrane domain. In particular, annexin A2 forms heterotetramers at the plasma membrane with its ligand p11, and it is enriched at the apical domain through its association with PtdIns(4,5)p2 (Martin-Belmonte et al., 2007; Rescher et al., 2004). The delivery of sucrase-isomaltase to the apical domain in Madin–Darby Canine Kidney (MDCK) cells requires annexin A2 ( Jacob et al., 2004). Furthermore, recent results have shown that Annexin A2 is required for the formation of the apical membrane and the lumen in MDCK cysts in a pathway that also involved Cdc42 (Martin-Belmonte et al., 2007).

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As mentioned above, the last step of fusion of vesicles with a target plasma membrane is mediated by the SNARE complex, denominated v-SNAREs, and t-SNAREs on the vesicle and target membrane, respectively. In polarized epithelial cells, apical and basolateral vesicles contain different v-SNAREs (such as TI-VAMP for apical, VAMP8 and cellubrevin for basolateral transport).The t-SNAREs are localized to the apical (syntaxin-3 and SNAP23), and basolateral (syntaxin-4 and SNAP23) plasma membranes. Loss-of-function or mislocalization of SNAREs leads to a concomitant disruption of plasma membrane delivery of the apical or basolateral vesicle population (ter Beest et al., 2005).

2.2. Cell–cell junctions and polarity complexes Epithelial polarity in multicellular organisms is also regulated by the formation of cell–cell junctions between cells, as well as by the presence of polarity complexes conserved throughout evolution. The polarity program is partially executed in epithelial cells by the competition and sequestration of these polarity complexes in different subcellular domains. 2.2.1. Junctional complexes All along the lateral membrane that connects neighboring cells, the apical– basal axis is delimited by junctional complexes. In vertebrates, these complexes include apical tight junctions (TJs), laterally localized adherent junctions (AJs), desmosomes and the connect cells with the ECM. Analogous structures exist in the Drosophila epithelium with some small differences. The Drosophila equivalents of the TJ, known as the septate junction, are localized basal, rather than apical to the AJ, although their cellular functions appear to be similar. For the purpose of this review, we will concentrate our discussion on vertebrate TJs and AJs. TJs serve not only to establish an apical–basal barrier that inhibits the diffusion of solutes across the epithelial layer (gate function), but they also restrict the movement of proteins and outer-leaflet lipids between the apical and the basolateral membranes (fence function) (Matter and Balda, 2003). The TJs are composed primarily of the membrane-bound junctional adhesion molecules ( JAMs), claudins and occludins, which are connected to the cytoskeleton through the PDZ containing proteins zonula occludens 1–3 (ZO-1, ZO-2, and ZO-3). These adapters also recruit regulatory proteins, such as protein kinases, phosphatases, small GTPases, and transcription factors to the TJs. This protein scaffolding facilitates the assembly of highly ordered structures that regulate epithelial cell polarity, proliferation, and differentiation (Kohler and Zahraoui, 2005). AJs perform multiple functions including initiation and stabilization of cell–cell adhesion, regulation of the actin cytoskeleton, intracellular signaling, and transcriptional regulation. The central part of the AJs includes

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interactions among transmembrane glycoproteins of the classical cadherin superfamily, such as E-cadherin, and the catenin family members including p120-catenin, b-catenin, and a-catenin. Together, these proteins control the formation, maintenance, and function of AJs (Hartsock and Nelson, 2008). E-cadherin is the major transmembrane protein of the epithelial AJs, and initiates intercellular contacts through trans-pairing between cadherins on opposing cells (Gumbiner, 2005). Classical cadherins also bind directly and indirectly to many cytoplasmic proteins, particularly members of the catenin family, which locally regulate the organization of the actin cytoskeleton, cadherin stability, and intracellular signaling pathways that control gene transcription (Perez-Moreno and Fuchs, 2006). Formation of the AJs leads to assembly of the TJs, but E-cadherin is not required to maintain TJs organization (Capaldo and Macara, 2007). Recent results have shown that an hepatic cell line unable to form AJs can form functional TJs in a delayed manner (Theard et al., 2007). 2.2.2. Polarity complexes Throughout evolution and the development of highly complex multicellular organisms, the requirement for cell asymmetry has remained essential. Given this, it is not surprising that many of the protein networks that have evolved to regulate cell polarity in Caenorhabditis elegans, Drosophila melanogaster are conserved through all eukaryotic species. In fact, each protein present in the polarity complexes involved in regulating cell polarity in C. elegans and Drosophila has highly conserved mammalian homologues. Over the past 15 years, the identification and analysis of either single or multiple mammalian homologues of the Drosophila polarity proteins have served to validate a conserved functional role for these proteins in regulating epithelial cell polarity. Furthermore, the manipulation in cultured mammalian cells has allowed us to gain new insights into the physical and functional interactions of the polarity proteins, and the coordinated development of apical– basal polarity in epithelium that was not possible in other in vivo models. Par4, also called LKB1 in mammalians, is mutated in Peutz–Jeghers syndrome (PJS), causing a predisposition to benign and malignant epithelial tumors (discussed in Chapter 5 of this volume). Activation of LKB1 by STRAD in mammalian intestinal epithelial cells in culture has demonstrated an essential role in cell polarity: cells with activated Par4 become polarized in the absence of cell–cell and cell–extracellular-matrix interactions, form an actin-rich apical brush border, and localize the TJs component ZO-1, and the AJs component p120 around it (Baas et al., 2004). LKB1 phosphorylates many kinases, including both the polarity protein, Par1 (Fig. 3.3) and AMPK. One particularly interesting finding is the involvement of LKB1 and AMP Kinase (AMPK) as master regulators of the energy state of the cell. During energy stress, LKB1 and AMPK are vital for epithelial polarization in both Drosophila and cultured mammalian cells, but apparently, they are

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Figure 3.3 The signaling pathways regulated by the polarity complexes for epithelial polarity in mammalian cells. The acquisition of apical–basolateral polarity during epithelial morphogenesis requires the activity of several polarity complexes. Phosphorylation and binding of Par5 (black) provides a mechanism for mutual exclusion for different PAR proteins. The basolateral determinant Par1 (green) that has diffused onto the apical domain is phosphorylated by Par6/aPKC (white), which inhibits the Par1 kinase activity and induces binding to Par5, and release into the cytoplasm. The apical determinant Par3 (orange) that diffuses into the basolateral domain is phosphorylated by Par1, which induces binding of Par5 and release into the cytoplasm. STRAD (brown) phosphorylates LKB1 (light green), which in turn activates Par1. The basolateral determinant Lgl (light blue) that has diffused onto the apical domain is phosphorylated by aPKC and induces its release from the cell cortex into the cytoplasm, although the exact mechanism is still unkonwn. aPKC phosphorilates Crumbs (purple) at the apical cortex to control the extension of the apical surface and/or apical junction assembly, together with PatJ (grey) and Pals1 (light brown), forming the CRB complex. Lgl, Dlg (yellow) and scribble (light purple) form the SCRB complex to define the basolateral domain by inhibiting Par complex and the CRBs complex. The localization of Par6/aPKC to the apical cortex and TJ is mediated by activated Cdc42-GTP (blue) or Par3, respectively.

less necessary when the energy and ATP level of the cell is high (Lee et al., 2007; Mirouse et al., 2007; Zhang et al., 2006b; Zheng and Cantley, 2007). Par1 is a Ser/Thr kinase that is localized to the basolateral membrane and excludes apical determinants, such as Par3, from this domain (Fig. 3.3). In addition, Par1 determines the organization of microtubules in mammalian cells, which in turn establishes the position of the luminal surface (Cohen et al., 2004, 2007). High Par1 activity converts columnar epithelial cells with vertical microtubules and an apical luminal surface into a hepatic type of epithelial cells, with horizontal microtubules and lumens

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that form between adjacent cells. In addition, high Par1 activity also causes the direct transport of apical proteins to the apical domain be modified to transcytotic route (Cohen et al., 2004), and it might regulate the docking and fusion of vesicles at the plasma membrane, at least in yeast cells (Elbert et al., 2005). Apical–basolateral polarization is essentially mediated by three interacting protein complexes: (1) Par/aPKC complex, consisting of Par3 (Baz), Par6, and atypical protein kinase C (aPKC); (2) Crumbs complex, consisting of Crumbs, Stardust, PATJ and several recently identified components, such as Yurt and Tuberous sclerosis (TSC) (Laprise et al., 2006; Massey-Harroche et al., 2007); and (3) the Scribble complex, consisting of Scribble, Discs large (Dlg), and Lethal giant larvae (Lgl) (Fig. 3.3). These complexes have been reviewed in detail elsewhere (Goldstein and Macara, 2007; Wang and Margolis, 2007). The Par/aPKC complex is involved in polarity and spatial organization in almost all metazoan cells, whereas the Crumbs complex is more specific to epithelial cells. The functions of the Scrib complex in mammals include exocyst-dependent basolateral exocytosis, and apical junction regulation (mScrib inhibits junction formation at the basolateral domain) (Qin et al., 2005; Yamanaka and Ohno, 2008; Zhang et al., 2005). Even though the molecular nature of these relationships is mostly unknown, these three complexes interact by a system of mutual exclusion to define the apical and basolateral surfaces of epithelial cells in Drosophila (Bilder et al., 2003; Tanentzapf and Tepass, 2003), and they could function similarly in mammalian cells (Fig. 3.3). The Par/aPKC complex is a master regulator of polarity (Munro, 2006). Mammalian Par3 is localized to TJs through the interaction with JAM at the apical/lateral boundary (Izumi et al., 1998), and functions in their assembly (Chen and Macara, 2005), whereas Par6/aPKC maintains the integrity of the apical domain (Martin-Belmonte et al., 2007). Par6 acts as a targeting subunit for aPKC, and it recruits both Crumbs complex (Hurd et al., 2003b; Lemmers et al., 2004) and Lgl (Scribble complex) as substrates (Betschinger et al., 2005). CRB controls the extension of the apical membrane (Macara, 2004), whereas the Par3-mediated phosphorilation of Lgl restricts the localization of Lgl to the basolateral domain. On the other hand, the Scribble complex suppresses apical membrane identity in the basolateral domain by inhibiting the Par3 complex (Tanentzapf and Tepass, 2003). How these opposing activities lead to the coalescence of AJs into the mature zonula adherens is not known, although it could involve regulation of the polarized transport of membrane proteins. Lgl homologues interact with Myosin II (a regulator of the actin cytoskeleton), with components of the exocyst complex (which is involved in transporting proteins to the plasma membrane), and with the fragile X-associated protein FMR1 (which is involved in transport and translational control of specific mRNAs). Finally, recent data has implicated PKA in TJ assembly (Kohler and Zahraoui, 2005). PKA activity is required to recruit claudin1 and ZO1, and

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inhibits the association of vasodilator-stimulated protein (VASP), indicating that PKA signaling is important for the assembly of functional TJs. In addition, cAMP and PKA activity stimulate apically directed transcytosis and secretion in epithelial cells, and budding of constitutive transport vesicles from the TGN to the cell membrane (Kohler and Zahraoui, 2005).

3. Epithelial Morphogenesis into Tubes Tubular systems are generated using not only mechanisms for cell trafficking and polarization, but also mechanisms based on common elements of cell behavior associated with the three-dimensional (3D) architecture of the epithelial organs. Some of these mechanisms include cell migration to target sites, cell-fate diversification, and localization of specialized cells to different regions of the tube system. Tubular organs in vertebrates present a very complex organization with different cell types and localization (i.e., the kidney, the mammary, salivary and lachrymal glands). However, in simpler organisms, such as C. elegans and Drosophila, such epithelial organs consist of only few cell types, and a straightforward development program. In the past years many different cell models have been used to analyze the molecular and cellular events required to organize individual cells into 3D epithelial organs.

3.1. Model systems for studying tubulogenesis 3.1.1. In vitro models Several in vitro systems consisting on cultured epithelial cell lines grown in a layer of (or embedded in) extracellular matrix (ECM) have been developed to study the molecular and cellular events required to organize individual cells into epithelial organs. MDCK epithelial cell system is perhaps the best and most widely used in vitro model to investigate cell polarity during epithelial morphogenesis (Lubarsky and Krasnow, 2003; Martin-Belmonte and Mostov, 2008; Zegers et al., 2003). MDCK cells, which have properties of the kidney distal tubule and collecting duct, have been used for decades as a 2D model to study epithelial polarity and protein trafficking. However, since the filter support provides an overriding extrinsic cue to orient cell polarity, they represent a less appropriate model to analyze morphogenesis. By contrast, MDCK cells embedded in ECM form cysts, spherical monolayers enclosing a central fluid-filled lumen (Montesano et al., 1991), which have proven to be a very informative model system. Bissell, Brugge, and Werb labs have used 3D cultures of mammary cells for many years (Bissell et al., 2003; Mailleux et al., 2008; Sternlicht et al., 2006). Although, most mammary cell lines are not fully polarized, these 3D culture models illustrate how the microenvironment plays a critical role in regulating mammary

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tissue function and signaling. Experimental methods are provided to generate and manipulate 3D organotypic cultures to study the effect of matrix stiffness and matrix dimensionality on epithelial tissue morphology and signaling ( Johnson et al., 2007). Other in vitro systems use endothelial cells, a specialized form of epithelial cells, as a model (Bauer et al., 1992; Levin et al., 2007). The cell line EA hy926, derived from HUVECs cells, has proved to be a useful model for in vitro study of angiogenic processes. These cells plated on an ECM undergo a process of morphological reorganization leading to the formation of a complex network of cord, or tube-like structures. These events seem to resemble, in some aspects, an in vitro process of angiogenesis (Bauer et al., 1992). 3.1.2. In vivo models The most studied in vivo model systems for tubulogenesis include Drosophila trachea and salivary gland, zebrafish gut and vasculature, and single-cell kidney of C. elegans. The Drosophila trachea and salivary gland are genetic model systems for branched and unbranched tubes, respectively. Both organs begin as polarized epithelial placodes, which through coordinated cell shape changes, cell rearrangement, and cell migration form elongated tubes (Kerman et al., 2006). The last discoveries regarding the details of cell fate specification and tube formation in the two organs reveal significant conservation in the cellular and molecular events of tubulogenesis. In particular, in the morphogenesis of the Drosophila trachea, the control of cell invagination, migration, competition, and rearrangement is accompanied by the sequential secretion and resorption of proteins into the apical luminal space, a vital step in the elaboration of the trachea’s complex tubular networks (Affolter and Caussinus, 2008). A genetic approach using the zebrafish model has led to identification of mutations and molecules that are responsible for specification of endothelial cells, and differentiation of arterial and venous cells, as well as patterning of the dorsal aorta and intersegmental vessels. These studies highlight the unique utilities and benefits of the zebrafish system for studying development of embryonic blood vessels. In addition, the intestinal progenitor cells represent an excellent genetic model to analyze the complex process of intestinal morphogenesis, which involves interactions among multiple signaling pathways. Studies on morphogenesis are critical for elucidating the molecular basis of congenital gut defects and provide novel insights into intestinal oncogenic processes (Rubin, 2007; Zhong, 2005). Finally, although the nematode C. elegans has no kidney per se, it has proved to be an excellent model for studying renal-related issues, including tubulogenesis of the excretory canal, membrane transport and ion channel function, and human genetic diseases including autosomal dominant polycystic kidney disease (ADPKD) (Barr, 2005).

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Additionally, some labs have recently started to use ex vivo animal models (using mouse salivary and mammary glands mainly) that are generating abundant information about the molecular mechanism underlying tube formation and branching morphogenesis in superior vertebrates (Sternlicht et al., 2006).

3.2. Formation of epithelial tubes follows diverse patterns Strategies for making tubes can be classified according to whether the epithelial cells already have apical–basal polarity, or whether such polarity is acquired de novo in response to extracellular cues (Hogan and Kolodziej, 2002; Lubarsky and Krasnow, 2003). 3.2.1. Formation of tubes from already polarized epithelia Wrapping and budding are the two main mechanisms that have been described. Both require apical constriction, mediated by acto-myosin cables at the apex of the cells, and therefore are two mechanistically related processes (Lubarsky and Krasnow, 2003). Another common peculiarity is that in both mechanisms, tubes arise from a polarized epithelial sheet. The process for wrapping requires that a group of cells of an epithelial sheet undergoes apical constriction in a coordinated way, causing the bending of the epithelial sheet until the edges meets and seal to form a tubular structure with parallel orientation to the plane of the epithelium. Two examples of wrapping are the formation of the neural tube in vertebrates, and gastrulation in Drosophila (Dessaud et al., 2008; Leptin, 2005). The process for budding requires, similarly to wrapping, the invagination of the epithelial cells by apical constriction in a direction orthogonal to the epithelium plane, forming a new branched tube (Lubarsky and Krasnow, 2003). This mechanism is used during branching morphogenesis in many epithelial organs, such as salivary gland and the tracheal system in Drosophila (Kerman et al., 2006), as well as the formation of the mammalian lung (Metzger et al., 2008). 3.2.2. Formation of epithelial tubes from unpolarized groups of cells In a number of mechanisms described for epithelial tubulogenesis, tubes arise from clusters of cells or individual cells that are nonpolarized (Lubarsky and Krasnow, 2003). An important variable in these mechanisms for lumen formation is the requirement of cell death, which eliminates cells inside the luminal cavity. Experimental data obtained from different in vitro and in vivo models has characterized two basic mechanisms, cavitation and hollowing, for lumen clearance. In cavitation, the lumen is generated by apoptosis of cells in the middle of the structure, whereas in hollowing, the lumen is formed by membrane separation and/or repulsion. The mechanism of lumen formation in the 3D MDCK model can shift between hollowing

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and cavitation, depending on the degree of cell polarization (MartinBelmonte et al., 2008). However, the suppression of apoptosis by the expression of antiapoptotic factors, delays, but does not eliminate lumen formation, indicating that lumens eventually clear up by a caspase-independent death process, such as autophagy (Debnath et al., 2002; Mailleux et al., 2007).

4. De Novo Formation of Tubes: Conserved Pathways and Molecular Mechanisms Despite the diversity in the way cells assemble into tubes, there are many conserved morphogenetic processes. In this review, we have focused in summarizing the current findings obtained from the in vitro systems on the sequential steps implicated in the de novo formation of epithelial tubes. These include, at least, the orientation of the axis of polarity, and the symmetry breaking process at the level of the plasma membrane by the formation of the apical domain and the central lumen.

4.1. Initial cell–cell and cell–ECM interactions drive the formation of AJs and the orientation of the axis of polarity In a simple tube, all of the epithelial cells are oriented so that their apical surfaces face the central lumen (O’Brien et al., 2002). To build a tissue, the polarity of each cell must be coordinated. The first step for the organization of the epithelial architecture involves the concerted integration of polarizing cues from different sources. First, cells must sense their environment, including their position in relation to the surrounding cells. This can be mediated by direct interaction of cells with the ECM through a variety of receptors, such as integrins; and by the homotypic interactions of the cells with each other through cadherins to form cell junctions. The polarizing cues direct the cells on how they must organize spatially within the tissue, and how to orient the axis of polarity. To perform these functions, cells need to sense and modulate the interaction with the ECM; for instance, the stiffness and other mechanical and chemical properties of the ECM (Kass et al., 2007), and reorganize their cytoskeleton. The cadherin’s family of adhesion molecules controls the physical interactions between cells, and is particularly important for the dynamic regulation of adhesive contacts that is associated with diverse morphogenetic processes. In epithelial cells, E-cadherin forms the AJs, and facilitates the formation of the entire epithelial junctional complex (Gumbiner, 2005). In addition, during embryonic development, cadherins control the separation of distinct tissue layers, the formation of tissue boundaries, the changes in the shapes of tissues through cell rearrangements, and the conversions

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between different cell states (such as epithelium vs. mesenchyma) (Gumbiner, 2005). Furthermore, they are also essential in the maintenance of stable tissue organization, since they prevent the dissociation and spread of tumor cells (Cano et al., 2000). Cadherins associate with the actin cytoskeleton through the catenin family of proteins, and are associated with the cadherin-mediated cell adhesion at the AJs (Nelson, 2008). Although the association with the actin cytoskeleton is not required for the formation of the adhesive bond itself, it is required to produce the necessary force to generate the changes in cell shape and/or cell movement, to establish cell polarity and to organize the tissue. The orientation of epithelial polarity depends ultimately on the interaction of the cell with the surrounding ECM. Studies with epithelial cells grown in 3D, such as the 3D-MDCK system, have provided important information regarding how epithelial cells identify and interpret polarizing cues coming from the ECM, and how they respond by inducing signaling pathways that control the orientation of polarity. Rac1 was the first protein identified in this model to cause an inversion of polarity after disruption (O’Brien et al., 2001). Rac 1 (together with RhoA and Cdc42) belongs to the Rho-subfamily of small GTPases that function as chemical switches for biological processes (Etienne-Manneville and Hall, 2002; Jaffe and Hall, 2005). They cycle between GTP and GDP-bound states with different kinetics in specific subcellular compartments determined by their regulators: Rho GTP exchange factors (GEF) and Rho GTP activating proteins (GAP); and effectors (Bos et al., 2007). Further experiments demonstrated that the interaction of MDCK cells with collagen I causes activation of Rac1, since blockade of b1 integrin prevented activation of Rac1, and led to an inversion of orientation of cyst polarity, supporting the idea that Rac1 and b1 integrin are needed for normal orientation of polarity (Yu et al., 2005). Besides, activation of Rac1 induces polarized secretion and the assembly of laminin, which in turn activates Rac1 generating a positive feedback loop that reinforces the orientation signaling pathway (O’Brien et al., 2001; Yu et al., 2005). Recent findings have also demonstrated a role for RhoA, ROCK I, myosin II, PI3K, and protein kinase B in this polarity pathway (Liu et al., 2007b; Yu et al., 2008). How the orientation of epithelial polarity is determined is a key issue in biology. However, to differentiate this process from the establishment of polarity has been an arduous task for many years (Martin-Belmonte and Mostov, 2008; O’Brien et al., 2002). One reason is that in conventional epithelial cell culture experiments, cells are usually on an artificial substrate, which provides a strong cue for the cell to orient its apical surface pointing away from the support. An advantage of the 3D MDCK system is that the cells are in an isotropic environment lacking strong external cues to orient polarity and thus, are more sensitive to perturbations that alter the

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orientation of cell polarity. Additionally, a similar inversion of epithelial polarity has been detected in vivo during tumor progression (Adams et al., 2004). Interestingly, both integrins and cadherins seem to use similar signaling pathways to integrate the polarizing cues in the acquisition and maintenance of epithelial polarity. Previous work in vitro and in vivo has demonstrated important roles for small GTPsases in the establishment of the cadherin- and integrin-mediated adhesion. These include the Rho subfamily, mainly through Rac1 (O’Brien et al., 2001; Van Aelst and Symons, 2002; Yu et al., 2005), Arf GTPases and Rap1, a ras subfamily member (Fujita et al., 2006; Price et al., 2004; Rangarajan et al., 2003). In addition, integrins and cadherins have the ability to induce, through the PI3K, the generation of PtdIns(3,4,5)p3 (Kovacs et al., 2002; Velling et al., 2008), a phosphoinositide that controls the formation and identity of the basolateral plasma membrane (Gassama-Diagne et al., 2006) (discussed in detail below). Since GTPases and phosphoinositides regulate the cytoskeleton in different cell types and processes (Fukata et al., 2003), they could consequently mediate the connection of cadherins and integrins with the cellular cytoskeleton and other important processes associated with the formation of a polarized epithelia.

4.2. Vesicle trafficking, membrane separation, and lumen coalescence in lumen formation Once the epithelial cells have established apical–basolateral polarity and formed junctional complexes, the next step is to form the central lumen. The mechanisms by which this lumen is formed a key question in morphogenesis (Lubarsky and Krasnow, 2003). Emerging data from different models have concluded that, probably in all systems, epithelial cells create lumens following a series of events associated with membrane trafficking (Fig. 3.4) that include apical membrane biogenesis, transport of apical vesicles to the plasma membrane, secretion, and regulated expansion (Lubarsky and Krasnow, 2003; O’Brien et al., 2002). Lumen formation factors and other apical targeted proteins might be delivered to the nascent luminal surface by exocytosis of a specialized organelle, the vacuolar apical compartment (VAC), which is made of membranes that resemble the luminal plasma membrane. VACs are thought to be formed by endocytosis of a small portion of their external plasma membrane to create an internal vesicle containing some of the extracellular fluid (Fig. 3.4). These vacuoles fuse with each other and with the plasma membrane to produce small lumens between cells that expand to acquire the final size (Fig. 3.4). The existence of these VACs was characterized recently in vivo in the developing blood vessels of fish embryos (Kamei et al., 2006), although it could follow different morphogenesis programs (Blum et al., 2008). In epithelial cells, VACs seem to appear only in nonphysiological situations, either when the calcium levels are drastically

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Figure 3.4 Mechanism for de novo lumen formation in epithelia. Top, confocal sections of control and Cdc42-siRNA knock down MDCK cysts. Images of fixed cells in 3D were taken at the indicated times. Cysts are stained for an apical marker (gp135, red); adherent junction marker (b-catenin, green), and nuclei (blue). Arrowheads indicate accumulation of intracellular VACs close to the plasma membrane in MDCK cysts knock down for Cdc42. Scale bar 5 mm. Bottom, general mechanism for generation of lumens, including: (1) apical membrane biogenesis mediated by the endocytosis of apical proteins into VACs and vectorial transport of VACs to the plasma membrane, (2) fusion of VACs with the plasma membrane mediated by the Cdc42 dependent pathway, and (3) secretion, and regulated expansion.

reduced with chelators, such as EGTA (Vega-Salas et al., 1987), or when apical-vesicle delivery is delayed. In fact, the accumulation of apparent VACs was observed when Cdc42 was depleted in the 3D MDCK model, indicating a role for Cdc42 in the exocytosis of VACs to form the lumen (Martin-Belmonte et al., 2007). Confirming these observations, the expression of a dominant negative form of Cdc42, Cdc42N17, blocked capillary lumen formation in vitro (Bayless and Davis, 2002). Recent data has demonstrated a function for Cdc42 in the orientation of the mitotic spindle to position the apical surface during epithelial morphogenesis ( Jaffe et al., 2008), which suggest that Cdc42 must control different pathways during epithelial morphogenesis. In addition, other elements of the apical sorting machinery such as MAL/VIP17, Annexin 2, Annexin 13, Galectin-3, Syntaxin 3 and FAPP2, have been found to regulate lumen formation in the MDCK-3D model system (Martin-Belmonte et al., 2007; Torkko et al.,

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2008; Vieira et al., 2005). Rab11 and Rab8, two small GTPases implicated in membrane traffic, regulate exocytosis of apical proteins in vivo to form the lumen (Li et al., 2007; Sato et al., 2007). Epithelial secretion and endocytosis have also been required to drive airway maturation in the traqueal system (Behr et al., 2007; Tsarouhas et al., 2007). Finally, different proteins associated with vesicle transport in Drosophila photoreceptor cells have been characterized to have a role in lumen formation in morphogenesis (Beronja et al., 2005; Sang and Ready, 2002). A critical event for membrane separation and lumen formation is the presence of antiadhesive and/or repulsive factors in the apical surface to prevent the membrane from sticking. This may be mediated by secretion of large transmembrane glycoproteins or polysaccharides, which may induce membrane detachment by steric hindrance of cell–cell adhesion, and/or serve as a scaffold for the formation of the luminal space. For example, several groups showed that correct dilation of the tracheal tubes in Drosophila requires the formation of a transient luminal chitin-based matrix, which coordinates tube growth (Swanson and Beitel, 2006). Somewhat similarly, the Drosophila retina forms an epithelial lumen, the interrhabdomeral space, by the secretion of eyes shut, a protein closely related to the agrin and perlecan proteoglycan, into apical matrix of the luminal space. Eyes shut mutants fail to open the luminal cavity (Husain et al., 2006). In vertebrate development, renal glomerular epithelial cells (podocytes) undergo extensive morphologic changes necessary for creation of the glomerular filtration apparatus that include opening of intercellular urinary spaces. Podocalyxin, also known as gp135, a sialomucin, keeps the urinary space open by virtue of the physicochemical properties of its highly negatively charged ectodomain (Orlando et al., 2001; Takeda et al., 2000). This function for gp135/podocalyxin was confirmed recently in the 3D MDCK model (Meder et al., 2005). As mentioned before, membrane repulsion is a lumen formation event related to antiadhesive factors. Recent results have shown that Slit and its transmembrane receptor Robo play central roles in cardiac lumen morphogenesis functioning in autocrine signaling (Medioni et al., 2008; SantiagoMartinez et al., 2008). Slit–Robo signaling has an extensively studied function in repulsive neuronal axon guidance (Dickson and Gilestro, 2006). Why does lumen formation fail when Slit–Robo signaling is compromised? Medioni, Sanrtiago-Martinez, and co-workers have described that regulation of cell adhesion is a key factor, with Robo and E-cadherin having apparently opposing roles in lumen formation. Heart lumen formation is distinct from typical epithelial tubulogenesis mechanisms because the heart lumen is bounded by membranes that have basal rather than apical properties. In fact, lumens form as Slit–Robo signaling antagonizes E-cadherin-based adhesion specifically at the luminal domains of apposing cardioblast cells.

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4.3. The role of PtdIns in the definition of membrane identity Phosphoinositides have been implicated in nearly all aspects of cell physiology. Particularly, they function as general markers of membrane identity (Di Paolo and De Camilli, 2006). Diverse phosphoinositides are enriched in specific subcellular compartments, concentrated at the cytosolic surface of cellular membranes. The plasma membrane is enriched in phosphatidylinositol-4,5bisphosphate (PtdIns(4,5)p2), whereas PtdIns(3)p is found in endosomes and might specify the identity of various membranous compartments. Phosphatidylinositol-3,4,5-trisphosphate (PtdIns(3,4,5)p3) appears to be also associated with the plasma membrane and play conserved roles in the determination of cell polarity in very diverse cell types (Comer and Parent, 2007; Leslie et al., 2008). Furthermore, it has been proposed that the signaling mediated by (the balance of) PtdIns(4,5)p2 and PtdIns(3,4,5)p3 at the plasma membrane, together with the enzymes that regulate them, PTEN and PI3K, controls many of the most important cellular processes such as growth, polarity, motility, and proliferation (Carracedo and Pandolfi, 2008). PtdIns achieve direct signaling effects through the binding of their head groups to cytosolic domains of membrane proteins or cytosolic proteins. The PtdIns recruit the cytosolic proteins through phosphoinositide-binding modules, such as the pleckstrin homology domain (PH). Thus, they can regulate the function of integral membrane proteins, or recruit to the membrane cytoskeletal and signaling components (Di Paolo and De Camilli, 2006). 4.3.1. PtdIns 4,5 P2 is enriched at the apical domain and regulates the formation of the apical membrane and the lumen In mammalian epithelial cells, PtdIns(4,5)p2 is a key determinant of the apical surface, whereas PtdIns(3,4,5)p3 is a determinant of the basolateral surface (Gassama-Diagne et al., 2006; Martin-Belmonte and Mostov, 2007). In the 3D MDCK system, during the early stages of polarization, PtdIns(4,5)p2 becomes enriched at the apical membrane delimiting the newly formed lumen (Fig. 3.5). In contrast, PtdIns(3,4,5)p3 is exclusively localized to the basolateral membrane and excluded from the apical membrane (Fig. 3.5). The lipid phosphatase, PTEN (phosphatase and tensin homolog deleted on chromosome 10), which converts PtdIns(3,4,5)p3 to PtdIns(4,5)p2 becomes localized early to the apical domain, and its activity is required both for segregation of the two lipids, and for normal morphogenesis in different epithelia (Fig. 3.5) (Leslie et al., 2008). PtdIns(4,5)P2 functions for specific tethering of membrane binding proteins including cytoskeletal related proteins, such as vinculin, talin or ERMs, RhoGTPase activators, such as many Dbl family GEFs, and tethering complexes, such as annexin A2 complex and the exocyst (Liu et al., 2007a) and this may underlie the role of phosphoinositides in membrane identity. However, our understanding of how phosphoinositides are connected to cell polarity is still not complete.

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Figure 3.5 Cdc42 and phosphoinositides regulates the acquisition of apical– basolateral polarity during epithelial morphogenesis. PTEN activity mediates the enrichment of PtdIns(4,5)P2 (green) at the apical domain and restricts PtdIns(3,4,5)P3 (red) to the basolateral surface. The localization of PTEN to tight junctions is mediated by Par3, whereas its localization to the apical cortex could by mediated by NHERF. PI3K might localize to the adherent junctions (blue) and contribute to the stable presence of PtdIns(3,4,5)P3 at the basolateral membrane, which is necessary for the organization of this domain. PtdIns(4,5)P2, Annexin A2, and a specific GEF at the apical domain targets and activates Cdc42, which in turn activates Par6/aPKC and other effectors. Activated Cdc42 regulates the actin cytoskeleton (purple), which mediates the fusion of VACs with the plasma membrane to form the apical domain.

A possible explanation may involve the small GTPase Cdc42 and Par3. Cdc42 is a center molecule in different polarity processes in unicellular and multicellular organism (Etienne-Manneville, 2004). It is well known that PtdIns(3,4,5)P3 and PtdIns(4,5)P2 are required for the recruitment and activation of the Rho GTPases family to the plasma membrane, to modulate actin dynamics in various cell types (Etienne-Manneville and Hall, 2002; Yin and Janmey, 2003). For instance, targeting of PtdIns(4,5)P2 to the cell membrane leads to the recruitment and activation of Cdc42 in Xenopus eggs extracts. In cytoplasmic extracts from Xenopus eggs, PtdIns(4,5)P2 initiates an actin nucleation pathway that, synergistically with Cdc42, activates N-WASP, which in turn stimulates the actin-nucleating activity of the Arp2/3 complex (Rohatgi et al., 1999, 2000). In the 3D MDCK model, PtdIns(4,5)P2 induces the recruitment and activation of Cdc42 (MartinBelmonte et al., 2007). Cdc42 also binds to the Crib domain of Par6 and is necessary for correct localization of Par6/aPKC (Fig. 3.5), as well as for normal apico–basal polarization of Drosophila neuroblasts and epithelial cells (both in 3D MDCK and in Drosophila) (Atwood et al., 2007; Hutterer et al., 2004; Martin-Belmonte et al., 2007). Furthermore, recent

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results in Drosophila neuroblasts have shown that Cdc42 localizes Par6/ aPKC at the apical cortex in a Par3-dependent manner, indicating that Par3 is upstream of Cdc42 in these cells (Atwood et al., 2007), and upstream of Par6/aPKC in embryonic epithelial cells (Harris and Peifer, 2005). Par3 also plays an important role in connecting phosphoinositides and cell polarity. Par3 contains multiple PDZ domains and binds via its second PDZ domain to most species of phosphoinositides, including both PtdIns (4,5)P2 and PtdIns(3,4,5)P3 (Wu et al., 2007). This binding is needed for the recruitment of Par3 to the plasma membrane and for polarization of MDCK cells, at least in 2D culture. However, as Par3 binds to both of these lipids with nearly equal affinity, this lipid binding alone would not appear to explain the localization of Par3. In Drosophila epithelial cells, Bazooka/Par3 binds to PTEN and localizes it to the cell–cell junctions (von Stein et al., 2005). Although this interaction has also been described in vitro in MDCK cells (Fig. 3.5) (Wu et al., 2007), PTEN is also associated with the apical membrane in mammalian cells forming 3D structures (Martin-Belmonte et al., 2007). Therefore, more work will be needed to address what localizes PTEN in mammalian cells. Interestingly, PTEN interacts directly with the NHERF1 adaptor protein (Naþ/Hþ exchanger regulatory factor required to organize complexes at the apical membranes of polarized epithelial cells) through the PDZ motif of PTEN and the PDZ1 domain of NHERF1 (Morales et al., 2007). Additionally, Par3 malfunction has proved to induce defects in central lumen formation in vitro in the 3D-MDCK model system (Hurd et al., 2003a), and also in vivo in cardiac cyst development in mice (Hirose et al., 2006). 4.3.2. PtdIns 3,4,5 P3 is restricted to, and defines, the basolateral domain PtdIns(3,4,5)p3 is emerging as a spatial landmark of specialized regions of the plasma membrane, such as the axon growth cone, phagocytic cup, leading edge of chemotaxing cells and can even determine the orientation of the mitotic spindle (Toyoshima et al., 2007). Interestingly, addition of exogenous PtdIns(3,4,5)p3 to the apical surface of filter-grown MDCK cells causes the transformation of the apical surface into basolateral within a few minutes. These experiments show that phosphoinositides are indeed molecular determinants of the identity of the apical and basolateral surfaces even in 2D epithelial cultures. In further support of the hypothesis that PtdIns(3,4,5)p3 specifies the basolateral surface, growth of MDCK cells in the presence of low concentrations of inhibitors of phosphatidylinositol-3 kinase (PI3K) reduced the size of the lateral membrane and height of the cells. Remarkably, binding of Pseudomonas aeruginosa, a bacterial pathogen that infects epithelia through the basolateral surface, to the apical surface leads to localized accumulation of PI3K, PtdIns(3,4,5)p3, and basolateral proteins at the apical surface, similar to the experimental addition of PtdIns

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(3,4,5)p3 to the apical surface. In essence, the bacteria transform the apical surface into basolateral, thereby enabling invasion of the host cell (Kierbel et al., 2005, 2007). However, the molecular details that define these relationships in different cells and tissues are probably distinct. In Drosophila photoreceptor epithelial cells, the recruitment of PTEN to cell–cell junctions, spatially restricts PtdIns(3,4,5)p3 to a the rhabdomere, a highly specialized region of the apical domain of these cells (Pinal et al., 2006). This result highlights that PtdIns(3,4,5)p3 can have a flexible role in determining specialized regions of a wide variety of cell types. There are other situations where PtdIns(3,4,5)p3 is involved in remodeling a specialized domain of the cell surface. For instance, stimulation of kidney cells by insulin binding to the basolateral surface leads to accumulation of PtdIns(3,4,5)p3 at the apical surface, followed by insertion of an epithelial sodium channel into the apical membrane (Blazer-Yost et al., 2004). Together, PtdIns(4,5)p2 and PtdIns (3,4,5)p3 seem to play essential roles in epithelial polarity and plasma membrane identity.

5. Epithelial Polarity and Disease There is growing evidence that a wide array of mutations in epithelial polarity genes is implicated in human disease (both monogenic and polygenic). Some of these genes have house-keeping functions in many cell types and their disruption in knockout mice models results in embryonic lethality. Many are found mutated with altered structure or expression, and are cause of diverse pathologies. Interestingly, many epithelial genes are related to a higher risk of developing cancer, and some of them are well known proto-oncogenes and tumor suppressors. In this review, we first address the relationship between epithelial polarity and cancer, and then we focus on mutations that are involved in syndromes caused by trafficking alterations. Finally, we address cytoskeletal, phosphoinositide and GTPase related disorders.

5.1. Cell polarity and cancer A vast majority of malignant human cancers are originated in epithelial tissues. Epithelial neoplastic tissues are characterized by loss of cell–cell junctions, loss of epithelial apico-basal polarity, and an increase in proliferation rates. Cells evade apoptosis and cancer progresses, as they acquire the mesenchymal phenotype, invading the inner tissues, a process called EMT (Hanahan and Weinberg, 2000). The regulation of cell proliferation and its related genes have been extensively studied. However, the relationship between polarity and proliferation is just recently starting to be unveiled. Indeed, current evidence shows that many mutated genes identified in

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aggressive cancers, are related to cell polarity establishment and maintenance, suggesting that cell polarity could act as a noncanonical tumor suppressor (Lee and Vasioukhin, 2008). Cancer cells that have lost their polarity regulation are more prone to cause metastasis, which is the spreading of cancer cells throughout multiple body tissues, and a hallmark of advanced cancer (Dow and Humbert, 2007). The key factor in cell–cell junction formation in epithelia is E-cadherin (Takeichi, 1991). Most epithelial cancer cells either harbor mutations in E-cadherin genes or lose E-cadherin expression along the way towards EMT (Thiery and Sleeman, 2006). Induced loss of E-cadherin in intestinal tissues in mice either increase tumor migration or proliferation and mutations in CDH1 are normally found in stomach, pancreatic, and other epithelial cancers (Conacci-Sorrell et al., 2002; Perl et al., 1998). The Scribble and PAR complexes are both implicated in human cancer (Fig. 3.6). The members of the Scribble complex (Lgl, Scribble, and Dlg) are neoplastic tumor suppressors in Drosophila larvae, where their mutation causes overgrowth of imaginal disc cells and loss of cell polarity and tissue architecture, larvae dying before reaching pupation (Bilder, 2004; Bilder et al., 2000; Gateff, 1978; Peifer, 2000). Moreover, Lgl and Dlg loss causes metastasis when imaginal disc cells are injected in adult hosts (Woodhouse et al., 1998). In mammals, there are two homologous Lgl genes, Lgl1 and Lgl2. Lgl1 knockout mice show severe brain dysplasia and loss of polarity in neuroepithelial cells, but the expression of a homolog gene, Lgl2, probably provides a functional backup in the rest of tissues (Klezovitch et al., 2004). Mammalian Lgl, Dlg, and Scrib expression loss is commonly found in a wide array of epithelial cancers (lung, prostate, skin, breast and colon cancers, and Human Papillomavirus (HPV)-induced cervical cancers) (Humbert et al., 2003). Studies carried out using cells from tumour samples have shown that overexpression of Lgl and Dlg are capable of phenotype attenuation and restoring cell polarity (Kuphal et al., 2006; Massimi et al., 2004). In aggressive HPV strains, HPV 16 and 18, E6 oncoprotein is capable of targeting the Scrib complex for proteolytic degradation, a condition that correlates with invasiveness increase and progression of the cervical tumor (Massimi et al., 2004; Thomas et al., 2005). Finally, Lgl, Dlg, and Scrib mutations increase the metastatic behavior of oncogenic Ras-mediated cancers and Ras oncogenic activation increases E6-mediated transformation, synergizing along with Scrib and Dlg proteolytic downregulation (Fig. 3.6) (Brumby and Richardson, 2003; Storey and Banks, 1993). The PAR–aPKC complex is the main regulator of apico-basal polarity at the apical pole of the cell, where it phosphorylates and inactivates Lgl (Betschinger et al., 2003; Plant et al., 2003). In Drosophila, activation of aPKC promotes tumorigenesis in lgl mutants and constitutively active forms

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Figure 3.6 Epithelial cell polarity and cancer. Mechanisms and complexes implicated in epithelial cell polarity that promote tumorigenesis when become deregulated. PTEN mutations cause PIP3 accumulation, leading to upraised survival signaling via Akt activation. Annexin 2 (Anx2) expression is abnormal in breast, brain, pancreatic, and prostatic tumors. Downstream of Anx2 and Cdc42, PAR complex signaling defects are related to cancer through distinct manners. Atypical PKC (aPKC) is overexpressed in many human cancers, and constitutively active forms cause overgrowth in D. melanogaster imaginal discs. The Scrib complex proteins behave like tumor suppressors and are targeted for degradation by E6 oncoprotein, promoting Ras-mediated oncogenesis. Par3 is the target of oncogenic ErbB2, which disrupts tight junction signaling by inhibiting Par3 binding to the PAR complex. Tumor suppressor VHL protein regulates PAR complex activation through targeting Par6 for proteolytic degradation. Finally, oncogenic TGFb receptor mediates Par6 binding to Smurf1 E3 ubiquitin-ligase, inducing Rho GTPase degradation and depolimerization of the actin belt that secures polarized epithelial architecture, promoting EMT.

of aPKC cause epithelial polarity loss (Fig. 3.6) (Lee et al., 2006). Lgl mutant phenotypes can be rescued inactivating aPKC, suggesting aPKC deregulation could have a causal role in those phenotypes (Rolls et al., 2003). The main isoform of atypical PKC in mammals, aPKC-i has also been found amplified in human cancers (ovarian and nonsmall-cell lung cancers) and aPKC-i activity is needed for xenograft tumour development and anchorage-independent cell growth (Eder et al., 2005; Regala et al., 2005; Zhang et al., 2006a). Less studied aPKC-z is also hyperactivated in squamous-cell carcinoma of the head and neck, where it mediates EGF-induced MAPK activation, and in glioblastoma cell lines (Cohen

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et al., 2006; Donson et al., 2000). Par6 is a target for the serine kinase activity of TGFb-Receptor, which promotes EMT during both development and tumor progression (Fig. 3.6). Par6 phosphorylation results in Smurf1 E3ubiquitin ligase recruitment and specific degradation of RhoA at the apical pole, destabilizing the actin belt that provides the scaffold for apical junction complexes (Ozdamar et al., 2005). Another signaling receptor, ErbB2, overexpressed or amplified in the 25–30% of breast cancers, disrupts cell polarity and provides antiapoptotic tumour protection by displacing Par3 from the PAR complex (Fig. 3.6) (Aranda et al., 2006; Hynes and Lane, 2005). The von Hippel–Lindau protein (VHL) is an E3-ubiquitin ligase that is mutated in the hereditary clear cell renal cancer syndrome that carries the same name. VHL functions as a tumour suppressor, controlling aPKC activity through signaling Par6 for degradation, and its expression loss correlates with an increase in aPKC levels, affecting cell polarity and favoring cancer progression (Fig. 3.6). PTEN (Phosphatase and Tensin homolog deleted on chromosome 10) is another protein mutated in multiple epithelial cancers, implicated both in cell polarity and cell proliferation, and it is the disease gene in the hereditary Cowden cancer syndrome (Di Cristofano and Pandolfi, 2000). The main dysfunction associated to PTEN mutations is PI3K pathway deregulation, leading to sufficient and signal-independent cell growth and survival (Fig. 3.6) (Lemmon, 2008). As noted before, recent studies have related PTEN function with cell polarity development in MDCK cysts. The dual function of a tumor suppressor, acting both in cell polarity development and proliferation inhibition, is an important example of how both processes are intricately related. We can only speculate whether or not PTEN mutations promote cancer through its polarity-related functions. More studies using downstream effectors and the elucidation of a complete pathway for Cdc42 and Par6–aPKC apical activation remain to be key issues for answering these questions. PAR complex regulators and effectors are also partly responsible for effects in cancer disease. The main GTPase activator for the PAR complex, Cdc42, is frequently downregulated upon Ras hyperactivation, and loss of Cdc42 cooperates with oncogenic Ras in cells expressing Ras activated mutants, allowing tumorigenesis in Drosophila imaginal discs (Fig. 3.6) (Sahai et al., 2001). PAR complex effector PAR4/LKB1 is mutated in the heritable PJS (Hemminki et al., 1998). LKB1 mutations in PJS do not impair the kinase activity, but instead reduce the ability to promote cell polarization in gastrointestinal cells (Alessi et al., 2006). The recent discovery of AMPK as a cell polarity regulator (by means of actomyosin activation through rMLC phosphorylation) is another important example of the coordinated regulation that controls cell polarity and proliferation (Forcet and Billaud, 2007; Lee et al., 2007; Williams and Brenman, 2008).

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Annexins present an important role in human cancer progression. Most genome-wide cancer expression assays have pointed out at least some annexins as genes differentially regulated in neoplasia (Hayes and Moss, 2004). Prostate cancer cells downregulate Annexin A2 during cancer progression and Annexin A1 seem to have reduced level of expression in squamous carcinomas (Fig. 3.6) (Liu et al., 2003; Paweletz et al., 2000; Vishwanatha et al., 2004). In addition, Annexin A2 is overexpressed in pancreatic, breast and brain cancer, and its upregulation could be the cause of increased cancer angiogenesis to promote metastasis (Huang et al., 2008; Liu et al., 2003; Nygaard et al., 1998; Sharma et al., 2006; Vishwanatha et al., 1993). There is a constantly growing body of evidence supporting the cancer stem cell theory (Cho and Clarke, 2008; Clarke and Fuller, 2006). Stem cells are present in many adult tissues and represent a source for the renewal of epithelial cells in the skin, the gut, and the blood. This theory proposes that certain cells in the tumor behave like stem cells, and they could also be the key in the origin of cancer. Adult stem cells normally undergo asymmetric cell division, a process that allows them to provide differentiated progenitor cells while retaining their own proliferative capacity (Knoblich, 2008). The mechanism requires polarization of fate-determinants and is driven by cell polarity regulators, like the PAR and Scrib complex (Wodarz and Nathke, 2007). When asymmetric division goes wrong, stem cells divide into equivalent cells, both of which retain their renewal potential. Recent advances propose that this process could be at the heart of tumorigenesis (Morrison and Kimble, 2006; Wodarz and Gonzalez, 2006). Cancer stem cells (CSCs) have been isolated from many tumors in mammals (Dick, 2003; Lee et al., 2008; O’Brien et al., 2007; Singh et al., 2004; Wang and Dick, 2005). These immortal, undifferentiated, progenitors are somewhat similar to mutant Drosophila neuroblasts, which are unable to divide asymmetrically. Although mutations in most tumorigenic polarity genes in Drosophila do not cause any phenotype in most tissues in mammals, probably because of functionally redundant paralogues, there are well known cancer-related functions for many genes that regulate spindle orientation and the mitotic checkpoint. Aurora A and Polo are two kinases that are implicated in the spindle assembly checkpoint, and, when mutated or deleted, cells fail to segregate chromosomes correctly (Malumbres and Barbacid, 2007). As a result, daughter cells inherit unstable and abnormal genomes, where some genes may become deleted or amplified, predisposing for carcinogenesis. In higher eukaryotes, cells are embedded into three dimensional environments, with multiple mechanic and chemical interactions with their milieu, organizing with specific tissue architecture (Bissell et al., 2003). The maintenance of this microenvironment has proven to be essential to counteract the proliferative and invasive properties of cancers, as well as to

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keep cells differentiated (Yamada and Cukierman, 2007). Original studies in oncogenesis resulted from comparison between in vivo and in vitro tumorigenic cell lines. For example, Ras-mediated or oncovirus-mediated transformation is evident in cultured cells, but is insufficient to generate tumors by itself in a complete organism (Dolberg and Bissell, 1984; Frame and Balmain, 2000). Importantly, wild-type cells behaved in a completely different manner when they grew together with mutated cells in a tissue or in a culture plate. In culture, cancer cells seem to influence a tumor-like behavior on wild-type cells, while this is rarely seen in a complete organism where tumors do not normally change the phenotype of a whole tissue, but rather evolve clonally. In addition, some epithelial cancers behave differently in 2D versus 3D cultures. Single oncogenes that normally had effect in two dimensional cultures were unable to produce the same effect in 3D cultures (Partanen et al., 2007). A fundamental signal is provided to the cells through ECM interaction with integrins (Gumbiner, 1996). In mammary gland cells cultured in 3D, disruption of b1-integrin signaling (using blocking antibodies) was sufficient to attenuate ErbB2-mediated transformation and invasion (Weaver and Roskelley, 1997; Weaver et al., 1997). It was proposed that in mammary gland cancer integrin signaling could promote cell proliferation and invasion, through the Rac1 pathway and that it could protect epithelial cells from apoptotic cell death. Indeed, anoikis is a type of cell death associated with lack of ECM signaling (Zahir and Weaver, 2004). However, as stated earlier, integrin signaling in the 3D-MDCK model was proven essential for promoting adequate orientation of cell polarity (O’Brien et al., 2001). Recently, it was also proven that polarity could regulate apoptotic clearance of epithelial lumen, and that disruption of Cdc42 activated polarity signaling results in increase of apoptosis in developing cysts (Martin-Belmonte et al., 2008). We propose that apoptotic cell death could be activated by loss of polarity, as a tumor-suppressor mechanism that would prevent abnormal growth of deregulated cells, although the underlying mechanisms remain unclear.

5.2. Trafficking disorders As discussed previously, protein trafficking routes are among the most complex mechanisms in cellular physiology. Complex machinery including GTPases (Rab proteins), adaptor complexes (APs), and tethering and docking complexes (Annexins) have proven to be related to some extent with human disease (Fig. 3.7) (Aridor and Hannan, 2000, 2002). 5.2.1. Defects in protein sorting signals A wide number of monogenic diseases are caused by point gene mutations that distort the sorting signal, resulting in the incorrect distribution of a specific protein and originating the disease (Stein et al., 2002).

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Figure 3.7 Trafficking and cytoskeletal disorders. Defects in the correct sorting and trafficking of vesicles cause a wide array of human pathologies (in red). Alterations in peptidic sorting signals are responsible for CFTR mislocalization and defects in ion transport (Cystic Fibrosis, CF). Defective trafficking machinery (such as Myo5B or Rab8) originates dysfunctional apical membrane domains and apically targeted vesicle accumulation (Microvillus Inclusion Disease, MVID). Abnormally high levels of Rab activation are common in cancer disease, and defects in Rab targeting to vesicle membranes (caused by REP1 mutations) originate specific problems in specialized epithelia (such as retinal choroideremia). Vesicle formation is mediated by adaptor protein complexes, such as AP-3 which causes defects in lysosomal maturation in the Hermansky–Pudlack Syndrome (HPS). Finally, Rho family GTPases activation is mediated by Rho GEFs, which are overexpressed or gain-of-function mutated in many cancers (i.e., LARG, TIAM, and Dbl GEFs). Mutations in Rho GTPase effectors, such as Diaphanous 1 formin (Dia1) or WAS protein (WASP), inhibit actin polymerization and are cause for nonsyndromic deafness in humans and the Wiskott–Aldrich syndrome disease, respectively.

Cystic fibrosis (CF) is an autosomal recessive disease that affects chloride transport in epithelia and that arises through point mutations in a c-AMP sensitive chloride channel, CF transmembrane conductance regulator (CFTR) (Riordan, 2008). The protein is located to the apical membrane compartment in renal cells. Its cytoplasmic tail contains a PDZ binding motif that interacts with CFTR-associated ligand (CAL) (Moyer et al., 1999), and C-terminal apical trafficking motifs that interact with Naþ/Hþ exchange

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regulatory factor (NHERF) (Milewski et al., 2001). Mutations in the cytoplasmic tail result in loss of CFTR apical localization which in turn originates defective chloride transport and CF (Fig. 3.7) (Ameen et al., 2007). Recently, it was shown that myosin VI knock-out mice have a gastrointestinal CFTR phenotype, with loss of enterocytic brush border, implicating myosin VI in CFTR trafficking in vivo (Ameen and Apodaca, 2007). Wilson disease is a monogenic autosomal recessive disease of copper metabolism that affects hepatic copper excretion. The gene, ATP7B, codifies a cell surface receptor that forms complexes with copper and is transported apically to release the cation at the lumen of the epithelium (Forbes and Cox, 2000). High levels of copper regulate ATP7B trafficking to the apical membrane. ATP7B mutations affect either cation binding or ATP7B trafficking to late endosomes (LE), causing the liver to accumulate copper and ultimately producing metal toxicosis (Morgan et al., 2004). Familial hypercholesterolemia is an autosomal genetic disease caused by a failure in LDL removal from plasma by hepatocytes. The most famous LDLR mutation is the autosomal recessive mutation described by Brown and Goldstein, which alters LDLR clathrin mediated endocytosis (Brown and Goldstein, 1976; Davis et al., 1986). However, autosomal dominant familial hypercholesterolemia mutations also map to the LDL receptor gene (LDLR) and are of relevance to this review. LDLR is normally expressed and localized to the sinusoidal membrane through two tyrosine-based basolateral sorting signals (Matter et al., 1992). Point mutations in these signals translate into a missorting of LDLR to the apical membrane compartment and the inability of hepatocytes to eliminate LDL from plasma, leading to LDL accumulation (Koivisto et al., 2001). The consequences rise as heart disease and atherosclerosis because of LDL deposition, plaque formation, and inflammation at the arteries. 5.2.2. Defects in sorting signal recognition and vesicle formation—Adaptor protein complexes Deletion or disruption of either AP1 or AP2 is absolutely lethal and mutations are rare among population. While mammalian mutations in AP-4 have not been found, the model for a human disease, called Hermansky–Pudlak syndrome (HPS), resulted from mice which had mutations AP-3 subunits (Fig. 3.7), either in AP3D1 (Delta-adaptin) or in AP3B1 (b3A) (Feng et al., 1999). The AP3B1 mutations were finally discovered in a small number of human HPS patients (Dell’Angelica et al., 1999). The autosomal recessive syndrome is related to deficiency in protein sorting to lysosomal and lysosomal-derived organelles (such as melanosomes and platelet granules) and human patients suffer from oculocutaneous albinism and platelet storage disease, with variable symptoms arising in subphenotypes (HPS1-8). Other murine HPS genes have also been related with cell polarity and trafficking,

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such as Vps33a (which sorts proteins to the vacuole in yeast) and RabGGTase (the enzyme catalyzing geranyl-geranylation of RabGTPases) (Wei, 2006). 5.2.3. Defects in Rab GTPases RabGTPases regulate vesicle transport and fusion spatially and temporally in a large number of cellular pathways, which are normally orchestrated by a specific member of the family. Many Rab and Rab-associated proteins have been implicated in human disease, ranging from neuropathies to cancer (Seabra et al., 2002). Choroideremia is an X-linked retinal degeneration disease which ultimately leads to inoperable blindness. It is caused by mutations in the Rab escorting protein gene REP1 (Fig. 3.7) (Andres et al., 1993; Rak et al., 2004; Seabra et al., 1992). Another escorting protein, REP2 seems to be able to compensate REP1 deficiency in the rest of tissues, thus affecting only the retinal epithelium where both of them seem to be needed (Cremers et al., 1994). TSC is an autosomal dominant disease characterized by formation of hamartomas, renal failure, and mental retardation. Mutations in two proteins were discovered in TSC suffering patients and were named hamartin (TSC1) and tuberin (TSC2) (Crino et al., 2006). Both genes have tumor suppressor activity and their products form a complex that acts downstream of the insulin receptor signaling pathway in which they are phosphorylated and inactivated by Akt, halting mTOR signaling (Gao et al., 2002; Inoki et al., 2002; Potter et al., 2002). The complex has Rab GAP activity, specific for Rab5 and Rap1, implicated in endocytosis, and Rheb GAP activity, thus shutting down the mTOR pathway (Fig. 3.7) (Wienecke et al., 1995; Zhang et al., 2003). The complex also regulates polycystin-1 trafficking, and in patients with tuberin mutations, polycystin-1 mislocalizes to the Golgi, with consequent cystic disease (Kleymenova et al., 2001). Polycystin-1 is the most vastly mutated gene in renal ciliopathies (discussed later) and the fact that tuberin mutations impede its normal trafficking accounts at least in part for the renal phenotype of the disease. Not only disruption of RabGTPase function but also abnormal RabGTPase activation is an important cause of human disease (Fig. 3.7). Rab5a and Rab7 are overactivated in Thyroid autonomus adenomas, and Rab7 is also overexpressed in atherogenesis (Croizet-Berger et al., 2002; Kim et al., 2002). Rab1a, Rab4, and Rab6 are upregulated in Beta2-AR mutant mice (which are human cardiomyopathy models) (Wu et al., 2001). Rab25 is also amplified in ovarian and breast cancer cells and a novel prostate cancer mutated gene, PRC17 has shown to be able to effect as an upregulated Rab GAP for Rap1 and Rab5 in human prostate cancer (Caswell et al., 2007; Cheng et al., 2004; Pei et al., 2002).

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Rab8, a RabGTPase implicated in basolateral protein sorting and adherens junction assembly, has been related to retinal degeneration and intestinal microvillus inclusion disease (MID) (discussed later).

5.3. Cytoskeletal and phosphoinositide related disorders 5.3.1. Defects in Rho family GTPases and Rho regulators The role of GTPases in cancer has been reviewed extensively (Vega and Ridley, 2008). Many GTPase pathways are linked directly to cell proliferation, polarity and migration, and their alterations are intimately linked to cancer progression and metastasis. Results from in vivo GTPase studies are beginning to arise (Heasman and Ridley, 2008). Gene targeting and disruption in mice models have proven both Rac1 and Cdc42 knock-out mice are lethal and die during development, at embryonic stages. However, mutations and diseases related to RhoGTPase defects are usually linked to their regulators, GAPs and GEFs, and effector genes, which are more specific in function (temporally and spatially) and whose mutations do not affect a wide number of processes (Fig. 3.7) (Chen et al., 2000; Sugihara et al., 1998; Wang and Zheng, 2007). Some RhoGEFs are well characterized oncoproteins in vitro (Cerione and Zheng, 1996). Many activating mutations have been found in Dbl family of GEFs during analysis of tumour samples including LARG (leukemia-associated Rho GEF), TIAM (T cell lymphoma invasiveness and metastasis), and Dbl (Diffuse B-cell lymphoma). Dbl was originally isolated as an oncogene (diffuse B-cell lymphoma), while LARG was isolated as a LARG-MLL gene fusion in acute myeloid leukemia, and TIAM was found in a screening for invasiveness related genes using T cells. The TIAM specific activation of Rac and its upregulation and mislocalization in aggressive cancerous cells is the hallmark for the discovery of GTPase function in cancer metastasis and tissue colonization. Faciogenital dysplasia 1 (FGD1) is a Dbl family RhoGEF, specific for Cdc42 activation (Olson et al., 1996; Pasteris et al., 1994; Zheng et al., 1996). It is the disease gene for the hereditary Aarskog–Scott syndrome, characterized by facial and urogenital malformations. FGD1 is recruited to membranes where actin functions must be carried out, by means of its PH domain. Once there, it activates Cdc42 to regulate the cortical actin cytoskeleton. Typical mutations implicate gene translocation or nonsense mutations. However, a subset of point mutations affects the PH domain (Orrico et al., 2000). The phosphoinositide specificity of its PH domain has not been assayed, but this fact signifies the importance of GEF membrane-binding domains in vivo for efficient GTPase function and a role for PH domains in disease. WASP is the product of the gene mutated in Wiskott–Aldrich syndrome (WAS), a hematopoietic disease that is characterized by recurring infections (Fig. 3.7) (Aldrich et al., 1954; Derry et al., 1994). Insights into its function

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were provided by studying its homologue N-WASP, originally isolated from brain tissue. WASP is a Cdc42 effector and stimulates Arp2/3 dependent actin polymerization, supporting the cortical matrix, and the cellular morphology (Symons et al., 1996). WASP missense loss-of-function mutations are the most frequent cause of WAS, originating cytoskeletal abnormalities in hematopoietic linage cells and immune system defects. Another GTPase effector related with actin polymerization is Diaphanous 1 (DFNA1, Dia1), homolog of D. melanogaster gene (Lynch et al., 1997). Its autosomal dominant mutation causes nonsyndromic deafness in humans, an inner ear dysplasia (Fig. 3.7). Mammalian Dia1 (mDia1) belongs to the Diaphanous-related formin (DRF) superfamily (Goode and Eck, 2007). It is activated specifically by Rho binding, upon which it linearly polymerizes actin and works in concert with ROCK inducing stress fibers, and aligning actin and microtubules through its microtubule binding domain. The mutations affect the actin cytoskeleton of hair cells in the cochleosaccular region of the inner ear, causing morphological abnormalities and deafness (Muller and Littlewood-Evans, 2001). 5.3.2. Microvillus inclusion disease Microvillus inclusion disease (MID), or Davidson’s disease, is a familial enteropathy with autosomal recessive inheritance (Cutz et al., 1989). Enterocyte and colonocyte differentiation is impeded and cells develop abnormal epithelial polarity (Ruemmele et al., 2006). Intestinal cells loose their brush border apical structure, and apical proteins and microvilli appear in intracellular vacuoles (Ameen and Salas, 2000). It is an extremely rare disease (with only about 30 cases diagnosed since its discovery in 1978). The interest for the disease became of notice when the VACs were first proposed as intermediates in epithelial polarity development and lumen morphogenesis (see Section 4.2) (Vega-Salas et al., 1988). VACs resembled intracellular microvilliar structures observed in MID and suddenly it became clear that MID patients could carry mutations in some of the genes involved in VAC formation and/or exocytosis (Fig. 3.7). Initial experiments were directed towards the actin and microtubule cytoskeleton and cytoskeleton binding proteins. MID patients present decreased expression of actin, vinculin, and myosin. Moreover, disruption of actin or tubulin cytoskeleton using drugs and inhibitors proved to mimic the MID phenotype. However, the search for the MID gene proved to be extremely difficult, due to the low incidence of the disease. Recent observations in the field have shown that Rab8 knockout mice display the MID phenotype (Sato et al., 2007). Mutations in Rab8 have not been proven to correlate directly with MID in humans, but at least one patient shows reduced expression of the gene. Also, genetic studies in a MID family have revealed a series of mutations in MYO5B, a gene encoding a myosin V motor protein isoform. The mutations cause alterations in transferrin receptor trafficking and disrupt epithelial cell polarity, along with

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microvilliar inclusions (Muller et al., 2008). Interestingly, Rab 8 and myosin V interact during endosomal transport (Roland et al., 2007). These studies have demonstrated a crucial role for myosin V and Rab8 in basolateral and apical trafficking and their relationship with MID (Fig. 3.7). Although these are promising results, not all MID patients carry mutations in MYO5B or Rab8, indicating that the phenotype might be a common consequence of diverse alterations in apical trafficking machinery. 5.3.3. Defects in phosphoinositide metabolism As emphasized earlier, phosphoinositides are implicated in many aspects of polarized cell physiology. The enzymatic network that determines the concentrations of each phospholipid subspecies is under tight regulation, by means of diverse signaling routes, and results in accumulation of specific lipids in a specific temporal and spatial frame in the cell. Mutations in PTEN are perhaps the most well known and studied, because of their implication in cell transformation and cancer (discussed above). However, a wider role for phosphoinositide signaling in trafficking and human disease is starting to emerge (Halstead et al., 2005; Nicot and Laporte, 2008). Other mutations in phosphatases include SHIP1, SHIP2, myotubularins, and OCRL1. OCRL1 is the phosphatase that converts PtdIns(4,5)P2 into PtdIns(4)P, a fundamental signaling lipid for lysosomal sorting from the TGN (Lowe, 2005). Mutations in this gene are found in patients suffering the Oculocerebrorenal syndrome of Lowe. The syndrome affects epithelial lens cells, renal tubule cells, and brain cells, causing bilateral congenital cataracts, Fanconi syndrome, and neurodegenerative retardation. In kidney tubule cells, OCRL1 mutations cause PtdIns(4,5)P2 to accumulate at the TGN (where OCRL1 is normally localized, and probably exerting its function) (Choudhury et al., 2005; Zhang et al., 1998). As a result, lysosomal proteins are missorted to the apical region, affecting the polarized epithelial phenotype and normal tissue integrity, and highlighting the importance of phosphoinositides in membrane trafficking. However, as we explained before, PtdIns(4,5)P2 domains are also a platform for linking lipidic membranes to cytoskeleton. OCRL1 cells lose their ability to form stress fibers and other F-actin based structures, and F-actin regulators gelsolin and alfa-actinin are abnormally distributed. Also, OCRL1 is recruited to membrane ruffles by means of Rac activation in response to growth factor signaling. Recently, it has been found that OCRL1 interacts with diverse RabGTPases, including Rab1, Rab5, and Rab6 (Hyvola et al., 2006). Rab5 effector APPL1 recruits OCRL1 to clathrin coated vesicles during endocytosis through an ASH-RhoGAP-like domain present in OCRL1 (Erdmann et al., 2007). Some OCRL1 mutations locate in this domain and impede normal recruitment, affecting normal membrane to TGN retrograde transport (McCrea et al., 2008). Studies using Ocrl1 knockout mice have proven a functional overlapping between this gene and its homologue Inpp5b,

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explaining the absence of phenotype ( Janne et al., 1998). In summary, actin polymerization, lysosomal sorting, and receptor endocytosis all result from PIP2 metabolism impairment.

5.4. Cystic diseases of the kidney Cystic diseases of the kidney (CDKs) are a group of monogenic diseases characterized by the appearance of fluid-filled cysts and end-stage renal failure (Hildebrandt and Otto, 2005). ADPKD is the most frequent, occurring in 1 in 400 to –1 in 1000 live births and affecting about 5 million persons worldwide (Torres et al., 2007). Other CDKs such as autosomal recessive polycystic kidney disease, pleiotropic Bardet–Biedl syndrome, Alstrom syndrome, and Orofaciodigital syndrome type 1, are by contrast, very infrequent diseases. CDKs are among the most common genetic lethal diseases has made them one of the most interesting and most studied subjects in epithelial biology. Despite the decades of long effort, the disease mechanisms remain mostly unknown. Polycystic kidney cells are characterized by increased proliferation and apoptosis of collector duct epithelial cells and a renal concentrating defect in spite of aquaporin 2 and vasopressin V2 receptor expression. Positional cloning helped to identify the first two CDK genes, PKD1 and PKD2 that carried mutations in ADPKD type 1 or 2, respectively (Mochizuki et al., 1996). Localization studies proved that PKD1 and PKD2 proteins, polycystin-1 and polycystin-2, are widely expressed in epithelia, vascular smooth muscle, cardiac myocytes and other tissues, and are regulated through development. Both proteins are mainly present in the primary cilia in wildtype cells (Yoder et al., 2002). In particular, polycystin-1 is located at the plasma membrane and especially at the primary cilium, whereas polycystin2 is located mainly at the primary cilium and the endoplasmic reticulum but also at the basolateral plasma membrane. Cilia are membrane protrusions sustained over a tubular structure conformed by nine microtubule doublets, and originate from the basal bodies, a similar microtubular structure that is attached to the cellular centrosome. Nonmotile (nonflagellar) primary cilia are modular sensory organelles that allow vertebrate cells to receive chemical and physical stimuli of diverse nature. During cloning of other CDK genes and cystic phenotype related genes, it was found that their products localized at the primary cilium, basal bodies or centrosomes, or were involved in their biogenesis (Watnick and Germino, 2003). It has been proposed that both proteins form a complex that functions as a sensor for mechanic stress (Nauli et al., 2003). Polycystin-1 and polycystin-2 are large membrane glycoproteins (Hughes et al., 1995; Ward et al., 1996), with 11 and 6 transmembrane domains, respectively. Polycystin-1 binds other membrane proteins through its extracellular domains and polycystin-2 works as a calcium channel. When a mechanic stimulus, such as

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urine flow, pushes the primary cilium, protein interactions change polycystin-1 conformation and polycystin-2 increases calcium permeability. Increases in intracellular calcium effect as a second messenger and produce a series of changes in cell physiology (Clapham, 2007; Nowycky and Thomas, 2002). Mutations impairing the ciliary pathway would deplete cellular Ca2þ levels affecting the subsequent cascade of mechanisms (Rizzuto and Pozzan, 2003). Ca2þ levels regulate inversin recruitment at the centrosome (Simons et al., 2005). When calcium levels increase, inversin is released and targets Dishevelled (Dvl) for degradation, thus switching off Wnt signaling, a cellular pathway that promotes cell cycle through upregulation of G1-phase genes. Ca2þ levels also affect cAMP levels by inhibiting the main renal adenylate cyclase, AC6 (Borodinsky and Spitzer, 2006; Cooper et al., 1995). Furthermore, calcium inhibits Ras activation through CAPRI RasGAP, and also inhibits cAMP-mediated MAPK signaling, a pathway that activates transcription of proliferation genes and Orofaciodigital syndrome type 1 (Cullen and Lockyer, 2002). Ca2þ depletion due to polycystin complex dysfunction would cause Wnt, Ras, and MAPK signaling activation and cell proliferation, an increase in cAMP levels and PKA-mediated induction of proapoptotic genes and finally a general defect in Ca2þ signaling. The polycystin complex is also part of focal adhesion complexes during epithelial cell migration in renal development and localizes at cell–cell junctions in fully developed renal tubules (Huan and van Adelsberg, 1999; Kreidberg et al., 1996; Roitbak et al., 2004). The mechanosensory hypothesis concludes that the complex could act as a sensor receiving mechanic signals from the matrix during migration, from partner cells during epithelial differentiation, and from the lumen of the developing organ via the primary cilium. Supporting this hypothesis, certain mutations in polycystin-1 induce focal adhesion kinase (FAK) mislocalization and defects in focal adhesion signaling, and developmental polycystin dysfunctions cause embryonic cystic kidneys, a severe phenotype with bad prognosis.

6. Concluding Remarks The establishment and maintenance of cell polarity is a critical step for the development of the epithelial identity. Cell polarity requires the communication of epithelial cells between them and with the surrounding tissues and ECM. Understanding the molecular mechanisms that regulates cell polarity is critical to comprehend normal tissue homeostasis, as well as the development and progression of malignant diseases. Alterations in epithelial tissues are the cause of diseases that can affect from just few individuals every year, such as the MID, to millions worldwide, such as

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cancer. In fact, cancer was the direct cause of almost 8 millions of deaths in 2007, which represent 1/8 of the total deaths worldwide. In the next future, work in in vivo models and in mammalian 3D culture model systems will provide us with a clearer understanding of the mechanisms of polarity regulation during epithelial morphogenesis, and how to apply this knowledge in medical treatments.

ACKNOWLEDGMENTS We thank Carmen Martin Ruiz-Jarabo for comments on the manuscript, and members of the Miguel A. Alonso and Isabel Correas lab for discussion. Work supported by grants to FM-B, from the Ministerio de Ciencia e Innovacio´n (BFU2008-01916), the European Union (MIRG-CT-2007-209382) and HFSP (LT00426/2004-C). AER-F is recipient of a MS fellowship from Fundacio´n Obra Social La Caixa. An institutional grant from the Fundacio´n Ramo´n Areces to CBMSO is also acknowledged.

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C H A P T E R

F O U R

Functions of RAB and SNARE Proteins in Plant Life Chieko Saito* and Takashi Ueda† Contents 1. Introduction 2. General Features of Membrane Trafficking and Functions of RABs and SNAREs 2.1. RABs and SNAREs—How they work 2.2. New field of membrane trafficking research—RABs and SNAREs in plants 3. Molecular/Cellular Research on Plant RABs and SNAREs 3.1. The dawn of vesicular trafficking research in plants 3.2. Intracellular trafficking pathways involving RABs and SNAREs 4. RABs and SNAREs in Higher Order Plant Functions 4.1. Gravitropism 4.2. Tip growth 4.3. Autophagy 4.4. Plant–microbe interaction 4.5. Abiotic stress response 4.6. Cell differentiation, morphogenesis, and flowering 5. A Unique Regulatory System in Plants for Endocytic/Vacuolar Transport 6. Concluding Remarks and Future Issues Acknowledgments References

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Abstract In all eukaryotic cells, vesicular trafficking is crucial for maintaining cellular and organelle functions. RABs and SNAREs play key roles in vesicle/organelle identity and exchange. Budding yeast genetics and mammalian cell biochemistry were the most effective approaches for investigating molecular mechanisms underlying vesicular trafficking and remain important in exploring new horizons.

* {

Molecular Membrane Biology Laboratory, Advanced Science Institute, RIKEN, Saitama 351-0198, Japan Laboratory of Developmental Cell Biology, Department of Biological Sciences, Graduate School of Science, The University of Tokyo, Tokyo 113-0033, Japan

International Review of Cell and Molecular Biology, Volume 274 ISSN 1937-6448, DOI: 10.1016/S1937-6448(08)02004-2

#

2009 Elsevier Inc. All rights reserved.

183

184

Chieko Saito and Takashi Ueda

The field of vesicular trafficking attracted plant biologists in the early 1990s. Today, this field continues to stimulate a wide range of research. This review starts with some history of RAB and SNARE research in yeast and mammals and introduces a widely accepted general model. Then we summarize recent reports regarding plant RABs and SNAREs, focusing on functional diversity. Finally, we discuss how plants assign new roles to conserved vesicular trafficking proteins to perform divergent, higher order plant functions. Plants have apparently evolved a unique set of plant-specific RAB and SNARE molecules that play significant roles in plant life.

1. Introduction Eukaryotic cells have a sophisticated system of intracellular, single membrane-bound structures called organelles (Fig. 4.1). Each organelle is a compartment defined by a membrane with a specific composition of proteins and lipids (Warren and Mellman, 2006). The establishment and maintenance of compartmentalization depends primarily on coordinated vesicular

Plasma membrane

RAB Secretory vesicle

Arf RAB Endosome

Arf

Sar/Arf family Rab/Ypt family

Trans-Golgi network (TGN)

RAB Arf

RAB

RAB

Arf?

Golgi apparatus

Vacuole RAB

Nucleus

Sar

ER

Figure 4.1 Small GTPases that function in membrane trafficking in plant cells. Green and orange. Arrows indicate the pathways involving the Rab/Ytp and Sar/Arf families, respectively. The broken blue arrow indicates the recently proposed early endosomal property of the TGN.

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trafficking. ‘‘Membrane trafficking’’ is also a well accepted synonym for ‘‘vesicular trafficking’’ because transport vesicles vary from ‘‘small spherical vesicles’’ to ‘‘larger irregularly shaped vesicles’’ or ‘‘fragments of a donor compartment.’’ For simplicity, in this review, we shall use the term ‘‘vesicular trafficking.’’ Vesicular trafficking of soluble cargo and membrane proteins begins with the budding of transport vesicles from donor membranes and ends with their fusion to target organelles (Fig. 4.2). Strictly regulated vesicular transport between donor and acceptor compartments enables each compartment to have different sets of proteins inside and in the boundary membranes. This compartmentalization permits the maintenance of specialized environments required for the various chemical reactions important in cellular activities (Warren and Mellman, 2006). A single round in the transport process (from vesicle budding to its fusion) basically involves two classes of Ras-like, small GTPases known as SAR/ARF and RAB (Figs. 4.1 and 4.2). SAR/ARFs act on the budding vesicle while RABs function in targeting and/or tethering transport vesicles to acceptor compartments (Fig. 4.2). The number of RAB genes is generally much larger than the number of SAR/ARF genes. For example, there are a dozen RAB/Ypt (yeast protein two) GTPases and only one SAR GTPase and three ARF GTPases in the budding yeast, Saccharomyces cerevisiae. Table 4.1 summarizes the numbers of RAB found in S. cerevisiae, (budding yeast), Schizosaccharomyces pombe (fission yeast), Caenorhabditis elegans (worm), Drosophila melanogaster (fruit fly), Arabidopsis thaliana (plant), and Homo sapiens (human). About 60 members of the RAB/ Ypt family have been identified in humans and A. thaliana, the model plant. It is believed that each individual step in the vesicular transport pathway requires at least one RAB/Ypt GTPase. Thus, the high number of RAB

Budding

Sar1

ER

Fusion

Transport vesicle

RAB (Ypt)

cis-Golgi

Figure 4.2 Two major groups of small GTPases, SAR and RAB, act as molecular switches to regulate membrane trafficking. SAR/ARF facilitates vesicle budding from donor organelles, and Ypt/RAB facilitates vesicle tethering/fusion to target membranes. The ER–Golgi pathway is depicted in this figure as an example.

Table 4.1 Number of members of vesicle-trafficking protein families –

S. cerevisiae

S. pombe

C. elegans

D. melanogaster

H. sapiens

A. thaliana

Coat complexes (subunits) RABs SNAREs Qa-SNAREs Qb-SNAREs Qc-SNAREs R-SNAREs Sec1 Total predicted genes

6(31)

6(31)

6(29)

6(29)

7(53)

8(59)

11 21 7 5 6 5 4 6241

7 – – – – – – 6241

29 23 9 7 4 6 6 18,242

26–29 20 7 5 5 5 5 13,601

60 35 12 9 8 9 7 30,000–50,000

57 66 19 12 14 18 6 27,000

RAB and SNARE in Plants

187

GTPases in humans and plants indicates these multicellular organisms have more complex systems of internal membranous organelles than those in unicellular organisms, including S. cerevisiae or S. pombe. Membrane fusion is the final step in vesicular trafficking and requires another family of membrane proteins known as SNAREs. The number of SNAREs is also high in multicellular organisms (Table 4.1), and the proliferation of some SNARE subgroups is especially significant in land plants including A. thaliana. These observations raise the following questions: (1) why do plants have a plethora of RABs and SNAREs? (2) how are these proteins differentiated functionally? (3) what roles do RABs and SNAREs play in plant life? In this review, we first summarize the general functions of RABs and SNAREs that have been identified and characterized in yeast or mammalian systems (Section 2.1). We then provide an overview of plant research (Sections 2.2 to 5), from the pioneering work in the early 1990s to recent progress achieved in twenty-first century.

2. General Features of Membrane Trafficking and Functions of RABs and SNAREs The RAB GTPase was one of the first molecules found to be involved in the regulation of vesicular trafficking. It was first identified as a Ypt in S. cerevisiae, which was later shown to function in the secretory pathway (Gallwitz et al., 1983; Goud et al., 1990; Schmitt et al., 1986; Segev and Botstein, 1987; Segev et al., 1988). The mammalian counterparts were identified by screening rat brain cDNA libraries, and thus, they were termed RAB (‘‘rat brain’’) proteins (Touchot et al., 1987). In this review, we shall use the term ‘‘RAB’’ for proteins in the RAB/Ypt family. SNARE proteins were initially identified as components of synaptic vesicles, secretory vesicles that mediate neurotransmitter release at neuronal synapses. SNAREs were purified on the basis of their ability to form complexes with the soluble N-ethyl-maleimide-sensitive factor attachment protein (SNAP) (Clary et al., 1990). Thus, it was named the SNAP receptor (SNARE). SNAREs fasten transport vesicles to acceptor compartments to facilitate fusion. As will be described below, the specific targeting of vesicles is accomplished by RABs and tethering molecules (tethers). Then, a specific combination of four cognate SNARE molecules assemble to form a tight complex that brings the donor and acceptor membranes more closely together to facilitate membrane fusion.

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2.1. RABs and SNAREs—How they work 2.1.1. The GTPase cycle RAB GTPases comprise the largest subfamily of the Ras superfamily of proteins. Each type of RAB has a characteristic distribution on organelle membranes, where it regulates fusion events. In turn, each organelle has at least one type of RAB protein on its cytosolic surface. Figure 4.3 summarizes the subcellular localization of different types of RAB GTPases in mammalian cells. Like other small GTPases, RAB proteins cycle between an active GTPbound state and an inactive GDP-bound state. RAB also cycles between cell membranes and the cytosol, as illustrated in Fig. 4.4. The different nucleotide states function as a molecular switch; when activated in the GTPbound state, the membrane-bound RAB transmits signals to downstream effectors; in the inactive GDP-bound state, RAB detaches from the membrane. As RAB proteins have rather low intrinsic GTPase activity, they require help from the GTPase activating protein (GAP) to efficiently hydrolyze GTP. Once GTP is hydrolyzed, the GDP-bound RAB binds to the GDP dissociation inhibitor (GDI). GDI masks the geranylgeranyl moiety at the C-terminus of RAB, which is responsible for membrane association. Thus, GDI solubilizes RAB GTPases and holds them in cytosol until the next round of GTPase cycle. Activation of RAB is catalyzed by the guanine nucleotide exchange factor (GEF), which replaces the GDP with GTP on RAB proteins (Fig. 4.4). To execute signaling, RAB GTPases must detach from RAB GDI and travel to the appropriate organelle membranes. This step is facilitated by the GDI displacement factor (GDF) (DiracSvejstrup et al., 1997), which comprises a group of small membrane proteins that belong to the PRA/Yip family (Sivars et al., 2003).

Plasma membrane

Clathrin-coated pit

Plasma membrane RAB5

Secretory RAB8 granule

RAB4

Secretory RAB8 vesicle

trans-Golgi network

RAB5

STX16

Clathrin-coated vesicle

Gos28

Golgi apparatus

RAB7

Late endosome /Lysosome

STX5

cis-Golgi

Bet1

VAMP1

VAMP2

STX7

STX8

VAMP3

VAMP8

SNAP29 Membrin VAMP4

RAB9

ER

STX11

STX5 Golgi stack

Golgi stack RAB1 RAB6 cis-Golgi RAB1 RAB2

STX10

Eearly endosome

Eearly/ RAB8 Recycling trans-Golgi RAB8 RAB9 endosome RAB11 network

Golgi apparatus

STX5

STX5

Late endosome

STX7

STX8

VAMP3

VAMP8

STX7

STX8

STX18

Lysosome

STX18 Membrin

RAB1 ER

Figure 4.3 Subcellular localization of selected RAB (left) and SNARE (right) proteins in mammalian cells. Associated proteins are shown to the right of the structure names.

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RAB and SNARE in Plants

Nucleotide exchange

A

GEF GDP

GTP

GDP

GTP

RAB

RAB Inactive form (OFF)

Active form (ON) Pi GAP GTP hydrolysis

B Acceptor membrane RAB GAP GTP

RAB

GTP hydrolysis Pi RAB GDI GDP

RAB

RAB GDI

GTP

RAB

GDP

GTP

RAB

RAB

GTP

GDF

RAB GEF

Nucleotide exchange

GDP

Donor membrane

Figure 4.4 Model of RAB GTPase cycle. (A) RAB proteins cycle between the inactive (GDP-bound, green) and active (GTP-bound, magenta) forms. Two major classes of proteins are involved in this regulation. The RAB guanine nucleotide exchange factor (GEF, purple) exchanges bound GDP for GTP, and the RAB GTPase activating protein (GAP, brown) accelerates GTP hydrolysis on the active RAB protein.

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2.1.2. From GTPases to downstream effectors GTP-bound RABs bind to RAB effectors to evoke downstream reactions. A large number of RAB effectors has been identified by genetic and biochemical approaches (Cai et al., 2007; Markgraf et al., 2007; Novick et al., 2006). The RAB effectors can be categorized as (1) tethers, (2) regulatory factors (i.e., GEF or GAP), and (3) others. In this review, we will mainly discuss RABs from categories (1) and (2), which play important roles in vesicle targeting. Category (3) also contains very important RAB effectors, including APPL, a recently identified protein that directly links RAB5 to endosomal signaling. For more on Category (3) effectors, please see the original article and recent reviews (Benmerah, 2004; Horazdovsky, 2004; Miaczynska et al., 2004). Some RAB effector molecules are called ‘‘tethers’’ due to their function in attaching transport vesicles to target membranes. Tethers are divided into two groups: long fibrous proteins and large multiprotein complexes (Warren and Mellman, 2006). Table 4.2 summarizes selected RABs, GEFs, and tethers that function in the secretory and endocytic pathways in yeast, mammals (Markgraf et al., 2007), and plants. GM130, giantin, p115, and EEA1 are long fibrous proteins, while TRAPP, COG, exocyst, CORVET, and HOPS are multiprotein complexes. Below we will describe some examples of tether functions in endocytic and/or vacuolar/lysosomal trafficking. EEA1 is a long fibrous tether that targets endocytic vesicles to early endosomes and enables attachments between homotypic early endosomes (Fig. 4.5, left) (Warren and Mellman, 2006). The conserved FYVE finger domain in EEA1 binds PI(3)P, which is enriched in the endosomal membrane. This protein–lipid interaction anchors EEA1 to endosomes. In addition, the interaction between EEA1 and GTP-bound RAB5 is required for the attachment of EEA1 to early endosomes. The HOPS complex is a multiprotein complex involved in the homotypic fusion of vacuoles in yeast cells (Seals et al., 2000). HOPS consists of six proteins: Vps41p (Vam2p), Vam6p (Vps39p), Vps11p, Vps16p, Vps18p, and Vps33p (Fig. 4.5, right). The Vam6p component acts as a GEF for Ypt7p (yeast RAB7). The HOPS complex interacts with GTP-bound Ypt7p and vacuolar SNAREs (Sato et al., 2000; Stroupe et al., 2006) suggesting that this complex mediates the transition from tethering to

GTP, red; GDP, yellow. (B) RAB proteins cycle between the cytosol and various membranes. Once GTP is hydrolyzed to GDP by the action of RAB GAP, GDPbound RABs are detached from membranes and held in the cytosol by the RAB GDP dissociation inhibitor (RAB GDI, aqua). Prior to a new membrane fusion event, the GDI displacement factor (RAB GDF, blue) mediates RAB dissociation from RAB GDI. Then the RAB GEF facilitates the exchange of GDP for GTP, thus activating the RABs to evoke downstream functions via effector molecules.

Table 4.2 RABs, RAB GEFs, and Tethers in each trafficking step cis-Golgi –

Mammal

RAB RAB1 GTPase GEF TRAPP I

trans-Golgi/ trans-Golgi network Yeast

Plant

Mammal

Yeast

Plant

Mammal

Yeast

Plant

Ypt1

RAB1/ RABD ?

RAB11

Ypt31/32

–

Sec4

TRAPP II – Endosome Mammal RAB5

TRAPP II

RAB11/ RABA ?

–

Sec2

RAB8/ RABE ?

–

?

Yeast Ypt51/ 52/53

Plant RAB5/ RABF

Vps9 Class C

AtVPS9 ?

TRAPP I Tether p115 Uso1 – Early endosome Mammal Yeast RAB RAB5 Ypt51/ GTPase 52/53 GEF Tether

Rabex5 Rabaptin5 Rabenosyn5 EEA1 hVps34

Plasma membrane

Vps9 Vac1

? Plant RAB11/ RABA? – ? ?

hVps39* HOPS/ Class C

(CORVET?) Vps8 –

Exocyst Exocyst Lysosome/vacuole Mammal Yeast RAB7 Ypt7

hVps39 HOPS / Class C – –

Vps39 HOPS / Class C – –

? Plant RAB7/ RABG ? ?

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A

B

Vesicle

Cytosol

Cytosol Vacuole

PI3P ClassC

FYVE finger domain

41 Vam6

Tethering

Yp

EEA1

t7 Tethering

HOPS complex

RAB5

Vacuole

Early endosome

Figure 4.5 Two types of tethering factors. (A) Schematic structure of EEA1, an example of a long fibrous tethering molecule. Clathrin-coated vesicles are uncoated and tethered to early endosomes by EEA1. The FYVE finger domain of EEA1 binds to phosphatidylinositol-3-phosphate on the vesicle and the other end of EEA1 binds to RAB5 on the endosome. (B) Schematic structure of the HOPS complex, an example of a multiple protein complex tethering factor. The HOPS is a complex of cytosolic proteins that mediates tethering between homotypic vacuolar membranes.

SNARE-mediated membrane fusion. In mammalian cells, overexpression of the Vam6 counterpart leads to clustering of lysosomes; this supports a direct role for HOPS in membrane tethering (Caplan et al., 2001). Recently, a related tethering complex, CORVET, was identified; it contains four of the six components in HOPS (Peplowska et al., 2007). Interestingly, CORVET preferentially interacts with Vps21p (yeast RAB5) rather than Ypt7p, and the CORVET and HOPS complexes interconvert by dynamic subunit exchange. These lines of evidence suggest that tethering complexes are assembled modularly and mediate transitions from endosomes to lysosomes in cooperation with RAB GTPases. It was once thought that organelles were static compartments and small vesicles transported proteins and lipids between organelles. Recently, however, a new concept has emerged that sequential and directional membrane maturation occurs as a result of membrane trafficking. RAB GTPases are proposed to play important roles in the so-called RAB cascade (Fig. 4.6) (Cai et al., 2007; Markgraf et al., 2007; Novick et al., 2006). In this model, the effector of one RAB acts as the GEF for another RAB, which in turn recruits the next RAB to a maturing site to contribute to the directional maturation of membrane domains. The first RAB cascade was identified in the late step of the secretory pathway in yeast cells. Ypt31p and Ypt32p are redundant and highly similar RAB proteins. Through genetic and biochemical analyses, Sec2p, a known GEF for Sec4p, was also identified as an effector of Ypt32 (Fig. 4.6) (Ortiz et al., 2002). Similarly, the activating factor for Ypt32 was suggested to be an

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RAB and SNARE in Plants

A Ypt31/32

Sec2

c2

Ypt31/32

Vesicle

Sec2

c4

c4

Se

Se

t31

Yp

Se 32 1/ t3 Yp

/32 Sec2

Sec4

TGN

B EEA1 Rabenosyn 5

Early

Endosome membrane

Late

Figure 4.6 RAB-cascade and RAB conversion. (A) Yeast Ypt31/32 (purple) forms a RAB cascade with Sec2p (green) and Sec4p (red). The redundant yeast RAB GTPases, Ypt31p and Ypt32p, are involved in several post-Golgi trafficking steps. Sec2p is an effector of Ypt31/32p, and also functions as the GEF for Sec4p, another RAB GTPase. (B) RAB conversion from RAB5 (brown) to RAB7 (purple) might drive maturation of early endosomes to late endosomes. One of the RAB5 effectors, Vps11p, is a subunit of the conserved class C VPS/HOPS complex (light blue). Another subunit in this complex, Vps39p, displays GEF activity towards Ypt7p, a RAB7 ortholog in yeast. Thus RAB5, class C VPS/HOPS, and RAB7 appear to function in a sequential RAB cascade on endosomes. The gradual inactivation of RAB5 and activation of RAB7 are mediated by the HOPS complex. This replacement of RAB5 for RAB7 might cause maturation of early endosomes into late endosomes.

effector of Ypt1p, which regulates traffic between the endoplasmic reticulum (ER) and the Golgi (Wang and Ferro-Novick, 2002). The RAB cascade also seems to be an integral part of the endocytic pathway in mammalian cells. Rink et al. (2005) demonstrated that RAB5 on endosomes is displaced by RAB7 during endosomal maturation. The HOPS complex has been shown to play critical roles in this RAB conversion. A subunit of the human HOPS complex, hVps11, binds to GTP-bound RAB5 (Rink et al., 2005). Another component of the HOPS complex, Vam6p/Vps39p, has been demonstrated to be a GEF for Ypt7p, the yeast counterpart of RAB7 (Wurmser et al., 2000). Moreover, decreased activity of hVps39 was associated with delayed recruitment of RAB7 onto the progressively maturing endosome (Rink et al., 2005). These results strongly suggest that the RAB cascade, comprised of RAB5 and RAB7, plays an essential role in endosomal maturation in mammalian cells.

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In plant endosomes, two types of RAB5 proteins are reported to be localized in an overlapping manner (Ueda et al., 2004). This localization might suggest that two types of RAB5 mediate the maturation of plant endosomes. However, the mechanism and physiological significance of plant endosomal maturation remain undetermined. RAB effectors also play important roles in the compartmentalization of membrane domains, as demonstrated in studies of the Rabex5–RAB5– Rapabptin5 system (Fig. 4.7) (Grosshans et al., 2006). Rabex5 is a RAB5 GEF, which activates the initial recruitment of RAB5 to the membrane. The GTP–RAB5 binds to an effector, Rabaptin5, which further binds to Rabex5 to increase the GTP-exchange activity of Rabex5 on RAB5. These interactions generate a positive feedback loop that inhibits inactivation by GAP and GDI-mediated extraction from the membrane, thereby ensuring A Rab5 Rabex5 Rab5

Rabex5 Rabaptin5 Rab5

Endosome membrane

B

Rab5

Rabex5 Rabaptin5 Rab5

Rabex5 Rabaptin5 Rab5

Rabex5 Rabaptin5 Rab5

C EEA1 Rabex5 Rabaptin5 Rab5 Vps34

Rabex5 Rabaptin5 Rab5 Vps34

Rabex5 Rabaptin5 Rabenosyn 5 Rab5 Vps34

Figure 4.7 RAB5 GEF/effector complexes stabilize activated RAB5 on membranes to allow the compartmentalization of RAB5-positive membrane domains on endosomal membranes. (A) and (B) After settling onto the membrane of early endosomes, RAB5 is activated by its GEF, Rabex-5. GTP–RAB5 then interacts with its effector, Rabaptin5. RAB5, Rabaptin5, and Rabex-5 form a stable complex, which leads to an increase in the Rabex-5 exchange activity on RAB5, thereby maintaining RAB5 in its GTP-bound state. (C) The PI-3-OH kinase, VPS34, is a RAB5 effector recruited to endosomal membranes by Rabex-5/RAB5/Rabaptin5 complexes. VPS34 generates PI(3)P, which is consequently enriched in the early endosomal membranes. PI(3)P and GTP-RAB5 are both required to recruit other RAB5 effectors, including EEA1 and Rabenosyn5.

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RAB and SNARE in Plants

that Rab5 is maintained in the active state at subdomains on early endosomes. GTP–RAB5 further recruits other RAB5 effectors, including EEA1, that lead to downstream reactions like homotypic early endosomal fusion. 2.1.3. Tethering and fusion machinery As mentioned above, RABs and tethers bring transport vesicles in close proximity to target membranes. Subsequently, membrane fusion is mediated by SNARE molecules. Figure 4.3 summarizes the subcellular localization of SNARE molecules in mammalian cells. SNAREs are relatively small (
200–400 amino acids) proteins that are characterized by the presence of a particular helical domain called the SNARE domain ( Jahn and Scheller, 2006; Lipka et al., 2007). The SNARE domain is a stretch of 60–70 amino acids arranged in heptad repeats that have the propensity to form a coiled-coil structure. The majority of SNAREs possess one SNARE domain, but the SNAP-25-subtypes harbor two SNARE domains separated by a flexible linker. Most SNAREs associate with lipid bilayers via a C-terminal transmembrane (TM) domain (Fig. 4.8); however, some exceptions attach to membranes via a lipid anchor.

Donor membrane

Syntaxin13 (Qa)

Syntaxin6 (Qc)

VAMP4 (R)

Vti1a (Qb)

Acceptor membrane

Zwilling et al.(2007)

Figure 4.8 A modeled orientation of SNARE proteins in a SNARE complex (adapted from the graphic data (Zwilling et al., 2007) in the PDB database). The model is based on an X-ray structure of a SNARE complex composed of four helical cytosolic regions of SNARE proteins (Qa-syntaxin 13, Qb-Vti1a, Qc-syntaxin 6, and R-Vamp4). Their insertion into the membranes is inferred.

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Formerly, SNAREs were categorized into vesicle-associated (v-SNARE) and target membrane-associated (t-SNARE) groups on the basis of their localization. This categorization, however, cannot be applied to homotypic vesicle or organelle fusions that occur among endosomes and vacuoles. Alternatively, a classification based on the amino acid sequences in SNARE domain has currently been well-accepted (Fasshauer et al., 1998). Thus, depending on the conserved glutamine (Q) or arginine (R) residue in the center of the SNARE domain, SNAREs can be classified as Q- and R-SNAREs. In general, t-SNAREs correspond to Q-SNAREs and v-SNAREs correspond to R-SNAREs. Despite this new classification, vesicle resident R-SNAREs are often referred to as VAMPs (vesicle-associated membrane proteins). The Q-SNAREs are further divided into three subgroups, Qa-, Qb- and Qc-SNAREs, based on sequence similarity (Fig. 4.9). On the basis of their role in synaptic exocytosis, Qa-SNAREs are also frequently referred to as syntaxins. SNAP-25-like proteins comprise a special case because they harbor both Qb- and Qc- SNARE domains (Fig. 4.9; adapted from Lipka et al., 2007). Hereafter, we shall use the current terminology and refer to SNAREs as Q- and R-SNAREs. SNARE actions critically depend on the SNARE domains, which assemble into a tight cluster comprised of four coiled-coil helices. The four components of the SNARE complex are provided by one each of Qa-, Qb-, Qc- (or Qb þ Qc in the case of SNAP-25-like SNAREs), and R-SNAREs, and only complexes generated by correct combinations of cognate SNAREs can drive membrane fusion. In addition to the SNARE domain, the Qa-SNARE also contains an autoregulatory domain at the N-terminus (Fig. 4.9). This autoinhibitory domain consists of three helices known as the Habc motif in neuronal syntaxin. The Habc motif interacts with the SNARE domain in the same polypeptide. This intramolecular interaction is called the ‘‘closed’’ conformation (Fig. 4.9), and prevents the Qa-SNARE from assembling with other SNAREs. Sec1 family proteins could unfold closed Qa-SNAREs, thus converting Qa-SNAREs into the active form (also called the ‘‘open’’ conformation) and allowing complex formation (Burgoyne and Morgan, 2007; Toonen and Verhage, 2007). Plants also harbor a large number of RABs and SNAREs that diverge in a manner unique to plants. Recent studies have revealed incredibly interesting functions of these protein families, which we will describe in the next section.

2.2. New field of membrane trafficking research—RABs and SNAREs in plants 2.2.1. Life style of plants It is believed that single-celled organisms that lived on ancient Earth had already established many cellular functions that are still conserved in extant eukaryotes (Lloyd, 2006). Plants and animals separated from their common

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RAB and SNARE in Plants

N

A

Ha

Hc

Hb

Qa

TM

C Qa SNARE

N

Qb

TM

C Qb SNARE

Qc

TM

C Qc SNARE

N

N

C Qb + Qc SNARE

Qb

Qc

LD

R

TM

C R-SNARE

R

TM

C VAMP727

N

N

LD

B

Donor compartment

trans-complex

TM

Tethering Qa Hc

Closed

Qc Qb Qa

TM

LD

N

R

Ha Hb Hc Open

TM

Acceptor compartment

Component recycling Disassembly

Fusion and cargo release

cis-complex

Figure 4.9 Domain structure of plant SNARE subfamilies and a model for vesicle fusion via the SNARE complex. (A) General domain structure of plant SNARE proteins. Q-SNARE domains include the Qa(green), Qb(yellow), and Qc(red) domains. Plant R-SNARE proteins comprise R-SNARE domain (blue), the longin domain (orange), and the C-terminal transmembrane helices (TM). The seed plantspecific R-SNARE, VAMP727, contains a short amino acid insertion (blue section) in the longin domain. (B) A model for SNARE complex formation. Vesicle fusion is initiated by opening (activating) a closed Qa-SNARE on the target membrane.

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ancestor about 1.5 billion years ago and evolved independently into multicellular organisms. However, plants and animals still use the same initial tool set that they inherited from the common ancestor, including: genes, molecular machineries, and organelles. It is also predicted that the molecular framework of vesicular trafficking, including RAB and SNARE proteins, are well conserved among eukaryotes, because vesicular trafficking is a fundamental and essential cell activity. This prediction is partly correct, but recent studies have revealed that plants have evolved a unique mechanism of vesicular trafficking that fulfills plant-specific functions. There are significant differences between plants and animals at the cellular/organelle level. Plants have chloroplasts, large vacuoles, and rigid cell walls (Lloyd, 2006). Owing to chloroplasts, plants are autotrophic and need not move to find and catch food. In addition, large vacuoles occupy most of the total cell volume and a rigid cell wall cements cells together; these features are very important for land plants to survive under terrestrial environments, but also accentuate plant cell immobility. Through evolution, plants have acquired a unique mechanism for storing nutrients in seeds. These characteristics have conferred on plants a program for growth and development that is fundamentally different from animals, and gives plants the ability to thrive in a fluctuating environment. The biogenesis of vacuoles and a cell wall critically depends on vesicular trafficking, as is easily imagined, and recent studies have unveiled various higher order plant functions that owe much to an elaboration of vesicular trafficking. In the latter half of this review, we will focus on how plants have adapted the vesicular trafficking system to their life style. 2.2.2. Plants: The superb system for studies on vesicular trafficking From Mendel’s era, plants have been amenable to genetics. One of the most important reasons is that most plant species are hermaphrodites; a single flower or individual plant can produce both female and male gametes. Thus, together with C. elegans, plants offer a significant advantage over D. melanogaster or vertebrates for applying genetics. In the early stages of plant genetics, Zea mays (maize), Triticum aestivum (wheat), and Oryza sativa (rice) were chosen as experimental systems because of their agricultural importance. However, these plants were imperfect as a primary plant model, mainly because the large size of these plants and their genomes, and the rather long generation times made experimental design cumbersome. This exposes the Qa-SNARE domain and enables the assembly of a SNARE complex with helices from Qb-, Qc-, and R-SNAREs. R-SNARE is generally located on transport vesicles. Assembly into a trans SNARE complex causes an increase in the core a-helical structure density and then leads to transition to a cis complex, membrane fusion, and release of cargo into the accepter compartments (for details see Lipka et al., 2007; Jahn & Scheller 2006).

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In the 1980s, plant biologists selected a small dicotyledonous weed, A. thaliana, as their model organism. A. thaliana possesses excellent characteristics for a model plant; it has a small genome size (1.25 Mbp), a short generation time, and a small plant size. Nearly the complete genome sequence is currently accessible, and many research tools are available, including a large collection of T-DNA insertion mutants and a vast amount of genomic, proteomic, and functional data from previous analyses. Moreover, because this is a popular plant model, more data are accumulating every day. Another advantage offered by A. thaliana is that it is transformed easily and efficiently. A straightforward procedure using reverse genetics enabled large-scale T-DNA mutagenesis and expression of modified gene products, including dominant negative proteins and siRNA; these tools facilitated investigations into the physiological significance of genes of interest. Recent progress in the genome projects of other plant lineages has enabled comparative genomic analyses. The availability of new information and tools has dramatically accelerated studies on RABs and SNAREs in A. thaliana. Studies of RABs and SNAREs first performed in A. thaliana have now expanded to a variety of plants, including some crops. Vesicular trafficking has currently come into the limelight from a practical perspective. Many crops including rice, maize, wheat, and beans provide a major source of protein for humans and livestock. It is noteworthy that these crops accumulate storage proteins in subcellular organelles, including the ER and the vacuole. Thus, the transport, localization, and accumulation of many storage proteins depend on the vesicular trafficking system (Mu¨ntz, 1998). Furthermore, correct localization of many membrane proteins that are responsible for useful traits depends on the vesicular traffic (Ma et al., 2006, 2007; Miwa et al., 2007; Mu¨ntz, 1998). By manipulating the vesicular trafficking system, yield and nutrition can potentially be improved; this is an important aim of the research in this field. 2.2.3. RAB and SNARE families in plants Genome sequencing of various organisms, including S. cerevisiae, S. pombe, C. elegans, D. melanogaster, H. sapiens, and A. thaliana has revealed a divergent organization of RABs and SNAREs encoded in different genomes. Remarkably, plants have considerably more diversification in RABs and SNAREs compared with other eukaryotes, including subtypes unique to plants. The genome of A. thaliana contains 57 loci that encode RAB GTPases that have been phylogenetically grouped into eight clades (Bischoff et al., 1999; Pereira-Leal and Seabra, 2001; Rutherford and Moore, 2002). Six of the eight clades (RAB1, RAB5, RAB6, RAB7, RAB8, and RAB11) are related to RAB subclasses that are commonly conserved among plants, yeasts, and animals; the other two clades show high similarity to the mammalian RAB2 and RAB18 genes, which are not found in yeast. Among these clades, unique and significant divergence is found in the

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subfamilies RAB5, RAB6, RAB7, RAB8, and RAB11, which are all predicted to function in the post-Golgi trafficking pathway; that is, secretory and vacuolar/endosomal pathways (Rutherford and Moore, 2002). The genome of A. thaliana contains at most 66 loci encoding SNARE proteins (Lipka et al., 2007; Uemura et al., 2004; Yoshizawa et al., 2006). Comparative analysis of genome sequences among a wide range of green plant lineages, including Osteococcus tauri, O. lucimarinus, Chlamydomonas reinhardtii, Volvox carteri, Physcomitrella patens, A. thaliana, Populus trichocarpa, and O. sativa indicated that the SNAREs involved in post-Golgi trafficking pathways have also expanded and diverged in land plants. However, the number of SNAREs involved in the early secretory pathway shows only a little variation (Sanderfoot, 2007). The divergence associated with the postGolgi trafficking pathway suggests its importance in fulfilling plant-unique functions that were acquired during evolution.

3. Molecular/Cellular Research on Plant RABs and SNAREs 3.1. The dawn of vesicular trafficking research in plants It has been 25 years since the first identification of the yeast YPT1 gene in 1983 (Gallwitz et al., 1983). The first SNARE molecule was identified almost 20 years ago (Trimble et al., 1988), and 15 years ago, in 1993, the SNARE hypothesis was proposed (Sollner et al., 1993a,b). During the last decade, research on RABs and SNAREs has expanded and, accordingly, has received more and more attention. We will start this chapter with a little history on plant RABs and SNAREs. 3.1.1. The initial identification of RABs and SNAREs in plants 3.1.1.1. RAB Today, we know that plants do not harbor the RAS gene, but in the late 1980s many researchers tried to isolate the plant RAS gene, without success, by cross-hybridization using the mammalian RAS gene as a probe. Instead, some researchers identified another class of small GTPases in plants known as RAB (Anuntalabhochai et al., 1991; Matsui et al., 1989; Palme et al., 1992). Thus, the first plant RAB was called the A. thaliana ras-related protein (ARA; Matsui et al., 1989). Then, using plant RABs as probes, many other plant RAB GTPases were identified by screening various plant species, including A. thaliana (Anai et al., 1991), O. sativa (Youssefian et al., 1993), Nicotiana plumbaginifolia (Dallmann et al., 1992; Trimble et al., 1988), N. tabacum (Haizel et al., 1995), Z. mays (Palme et al., 1992), P. sativum (Drew et al., 1993; Nagano et al., 1993), Glycine max (Cheon et al., 1993), Vigna aconitifolia (Cheon et al., 1993), Brassica napus (Park et al., 1994), Beta vulgaris (Dallery et al., 1996), Medicago sativa ( Jonak et al., 1995), Petunia hybrida

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( Jako and Teyssendier de la Serve, 1996), Solanum lycopersicum (Loraine et al., 1996), Mangifera indica (Zainal et al., 1996), and Lotus japonicus (Borg et al., 1997). These genes were named inconsistently, according to the preference of the founders because there was no standard systematic nomenclature; this resulted in the current confusion in the names of plant RABs. Recently, a few nomenclatures have been proposed for A. thaliana RAB GTPases (PereiraLeal and Seabra, 2001; Zhang et al., 2007a), but these are not fully accepted because they are too complex and incomprehensible for people in other fields. Currently, the nomenclature issue remains under discussion, and a unified nomenclature will be announced in the near future. In this review, we will indicate RAB proteins by both the names designated in the original reports and the names proposed by Pereira-Leal and Seabra (2001), in addition, we will cite the names of the closest animal homologs for cross-referencing. 3.1.1.2. SNARE Plant SNAREs were first identified by either the ability to functionally complement yeast SNARE mutants or by homology to yeast SNAREs. These strategies resulted in the isolation of three vacuolar/endosomal SNAREs, AtPEP12 (now renamed SYP21), AtVAM3 (SYP22), and AtVti1 (AtVTI11) (Bassham et al., 1995; Sato et al., 1997; Zheng et al., 1999b). Several studies with forward genetic approaches have also identified some SNARE genes that were responsible for mutations associated with interesting phenotypes (Chapter 4 of this volume). The nomenclature for SNAREs was also disorganized initially; however, the naming system proposed by Sanderfoot et al. (2000) has now been generally accepted. Nevertheless, some SNAREs are still referred to with multiple names, reflecting the different ways they were identified (e.g., SYP111/ KNOLLE and VAM3/SYP22/SGR3).

3.1.2. Biochemistry, cytology, and forward genetics 3.1.2.1. RAB After the initial identification, each plant RAB underwent a basic characterization; its sequence was analyzed for similarities to mammalian or yeast RAB GTPases, its expression was profiled by Northern blotting, the number of homologous genes was estimated by Southern blotting, its GTP binding and GTP hydrolyzing activities were tested, and so on. Among A. thaliana RABs, the RHA1/RABF2a and ARA family members (ARA1–ARA5) were in the forefront in the early 1990s (Anai et al., 1991, 1994; Anuntalabhochai et al., 1991). One of the ARA members, ARA4/ RABA5C, was analyzed with immuno-detection and fractionation to determine its subcellular localization (Ueda et al., 1996b). However, information on subcellular localization of plant RABs was quite limited because specific antibodies against RABs were not widely available and only a few organelle markers were usable for determining subcellular localization. The functional implications of RABs in plant-unique functions was difficult to determine, but several intriguing results were reported.

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The sRAB1p, sRAB7p (G. max RAB1 and RAB7), and vRAB7p (V. aconitifolia RAB7) proteins were suggested to play critical roles in nodule formation (Cheon et al., 1993). The expression of PRA2 (a RAB11 homolog) in P. sativum was repressed by light, suggesting a function in stem growth mediated by phytochrome signaling (Nagano et al., 1993, 1995). RGP1 (a RAB11 homolog) in O. sativa was isolated by differential screening for genes whose expression was affected by 5-azacytidine treatment (Sano and Youssefian, 1991). Interestingly, the overexpression of RGP1 in tobacco plants caused an elevation of endogenous cytokinin levels, a hyperinduction of salicylic acid with wounds, and an increased resistance to a tobacco mosaic virus infection (Sano et al., 1994), though the molecular basis of these effects remains unknown. In addition to RAB GTPases, several RAB regulator molecules were identified in plants. The expression of ARA4/RABA5c in some yeast ypt mutants severely affected growth. By screening A. thaliana genes whose overexpression rescued the growth defect caused by RABA5c, a general regulatory factor for RABs was identified, a GDI (Ueda et al., 1996b). The same gene (AtGDI1) was also isolated by functional screening with a yeast GDI mutant, sec19 (Zarsky et al., 1997). A RAB escort protein (REP), believed to share the common ancestral molecule with RAB GDI, was found to be conserved in plants (Hala et al., 2005). AtVPS9a is the only currently known RAB GEF isolated from plants, and acts as an essential activating molecule for A. thaliana RAB5/RABF proteins (will be described later). Based on mammalian studies, plants should express a RAB GAP that plays an important role in the regulation of the GTPase cycle of RAB, but that discovery awaits future investigations. 3.1.2.2. SNARE Two A. thaliana SNAREs, AtPEP12/SYP21 and AtVAM3/SYP22, were isolated by functional screening using yeast mutants ( pep12 and vam3). Cell fractionation and immuno-electron microscopy revealed that they localized on prevacuolar compartments (PVC) and vacuoles, respectively (da Silva Conceicao et al., 1997; Sato et al., 1997). Two VTI1 homologs were subsequently identified in A. thaliana and were found to exert different effects on the yeast trafficking system (Zheng et al., 1999b). On the other hand, screening Xenopus laevis oocytes for molecules mediating abscisic acid (ABA) signaling identified a plasma membrane (PM) SNARE from tobacco, NtSyr1; this unveiled an intriguing link between membrane trafficking and hormone signaling (Leyman et al., 1999, 2000). Several SNARE genes were also found to be responsible for certain mutants in studies of forward genetics. This history is in contrast with that of the RABs: there were no examples of RAB genes identified through a forward genetic approach. KNOLLE was the first and best-characterized SNARE gene identified through forward genetics (Lukowitz et al., 1996). KNOLLE will be described in more detail later.

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3.1.3. Plant vesicular trafficking in the post genome era The completion of the A. thaliana genome sequencing (A.G.I., 2000) has brought an explosive increase of information. Owing to the development of microarray technology and the enrichment of various resources, we can now get an expression profile without Northern blotting or RT-PCR, and we can obtain mutants, marker cell lines, and cDNAs of interest by browsing a database and clicking the ‘‘place order’’ button. Live imaging technology has also grown to epoch proportions; fluorescent proteins, including green fluorescent protein (GFP) and its variants, provide great opportunities to visualize the subcellular localization of proteins in living cells. Moreover, in vivo fluorescent imaging is done with less time and labor compared to antibody-dependent cytological analysis. The advent of in vivo imaging has also drastically accelerated the accumulation of information on organelle dynamics and provided many tools for monitoring intracellular trafficking. These innovations have great impact on many areas in plant research, especially vesicular trafficking. The sequencing of other plant genomes followed that of A. thaliana and recently, the genome sequences have been completed for several green plants, including angiosperms, lycophyte, bryophyte, and green and red algae. Comparative genomics is an important research area today. It may unveil how and why plants have evolved plant-unique mechanisms for vesicular trafficking, and the significance of plant-specific RABs and SNAREs.

3.2. Intracellular trafficking pathways involving RABs and SNAREs In this section, we will overview each pathway of vesicular transport, focusing on the functions of RABs and SNAREs. An extensive analysis on the subcellular localization of A. thaliana SNARE was conducted (Uemura et al., 2004); thus, we can put most molecules into their putative places of function (Fig. 4.10). In this study, 54 SNARE molecules were tagged with GFP or Venus (a modified yellow fluorescent protein) and expressed transiently in protoplasts of A. thaliana suspension cultured cells. This method is very convenient and effective, but it should be kept in mind that overexpression in protoplasts sometimes causes the mislocalization of expressed proteins. Additional experiments testing the functions of tagged molecules under more physiological conditions should be conducted to ascertain the precise localization of each SNARE protein. The number of predicted SNARE molecules encoded in the A. thaliana genome varies in the literature (60 in Lipka, 2006; 65 in Yoshizawa, 2006; 64 in Sanderfoot, 2000). In addition to the 64 SNAREs described by Sanderfoot (2007), 2 putative SYP5 members were identified in A. thaliana

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Plasma membrane

SYP111

NPSN11

VAMP721

SYP112

NPSN12

VAMP722

SYP121

NPSN13

VAMP724

SYP122

VAMP725

SYP123

VAMP726

VTI12

SYP124 SYP125 SYP131

Endosome/ PVC/MVB

SYP132

VTI11 VTI11

VTI13

SYP51

SYP22

VTI14

SYP52

VTI13 SYP41

VTI11

SYP42

VTI12

SYP43

VTI13

trans-Golgi network

VAMP727

SYP21

VTI12

SYP61 Golgi apparatus

GOS11 GOS12

BS14a

SYP31

Mamb11

BS14b

SYP32

Mamb12

VAMP714

Vacuole

SYP51 SYP52 VAMP711

SYP71

ER

SYP81

SYP72

SEC22

SYP21

VTI11

VAMP712

SYP73

VAMP723

SYP22

VTI13

VAMP713

Figure 4.10 Membrane trafficking pathways in plant cells, where SNARE proteins of A. thaliana are localized. It should be noted that considerable numbers of SNAREs are localized on post-Golgi organelles. Qa-SNAREs are indicated in green, Qb-SNAREs in yellow, Qc-SNAREs in red, and R-SNAREs are in blue. Partial or weak localization is indicated by a lighter shade of color.

(Uemura et al., personal communication). Thus, the A. thaliana genome potentially encodes at least 66 SNAREs, but this requires further clarification to be conclusive. A considerable number of RABs in A. thaliana have been localized to distinctive organelles (Fig. 4.11). In addition, the similarity of RABs across diverse organisms allows us to predict the locations of some RAB subgroups. In this section, we attempted to map RABs and SNAREs in the network of vesicular transport pathways. We basically divide angiosperm RABs into eight clades (RAB1/RABD, RAB2/RABB, RAB5/RABF, RAB6/RABH, RAB7/RABG, RAB8/RABE, RAB11/RABA, and RAB18/RABC) according to Pereira-Leal and Seabra (2001) and

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Plasma membrane

Secretion RAB8 RABE

RAB11 RABA

RAB11 RABA

Cell plate Cytokinesis

TGN

RAB11 RABA

Vacuolar PVC Endosomal pathway RAB5 RABF RAB18 ? RABC

Golgi apparatus ER to Golgi Pathway RAB1 RABD

Golgi to ER Pathway

RAB6 RABH

RAB6 RABH

RAB2 RABB

RAB7 RABG

ER

Figure 4.11 Subcellular localization of plant RAB GTPases. RABs with predicted but unclarified locations are indicated by a lighter shade of color.

Rutherford and Moore (2002). We will indicate the plant from which each RAB was discovered with an ‘‘At’’ for A. thaliana, ‘‘Nt’’ for N. tabacum, and ‘‘Ps’’ for P. sativum in front of the acronyms for genes and proteins. 3.2.1. ER–Golgi 3.2.1.1. RABs 3.2.1.1.1. RAB1/RABD The RAB1/RABD group is widely conserved among eukaryotes. Extensive examinations in yeast and mammalian cells have revealed that the function of RAB1 members is to regulate traffic between the ER and the Golgi. The function of RAB1 homologs seems to be conserved in plants, because RAB1 homologs isolated from some plants were reported to complement a mutation in the yeast RAB1 ortholog, YPT1 (Kim et al., 1996; Park et al., 1994). Five RAB1 homologs encoded in the A. thaliana genome function in ER–Golgi trafficking, as demonstrated by the transient coexpression of mutant AtRABD2a (formerly called AtRAB1b) and fluorescent markers in tobacco cells (Batoko et al., 2000). The expression of a dominant negative mutant of AtRABD2a (N121I) resulted in the mislocalization of GFP markers that were targeted for secretion and the Golgi, but were retained in the ER. This effect was suppressed by the coexpression of wild-type AtRABD2a but not by AtRABE1d (formerly AtRAB8c), indicating a

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functional specificity for each RAB group. Intriguingly, the expression of AtRABD2a also caused the reduction or cessation of movement of the Golgi. These results suggest that RAB1 homologs are required for transport from the ER to the Golgi, and are somehow coupled with movement of the Golgi. 3.2.1.1.2. RAB2/RABB Budding yeast does not have a RAB2 homolog, but this RAB group is conserved in animals and plants. Mammalian RAB2 localizes on cis-Golgi membranes and acts in ER–Golgi trafficking. RAB2 is also reported to interact with Golgi matrix proteins. RAB2 homologs seem to be well conserved among green plants. Some members of the RAB2 group in A. thaliana and N. tabacum are expressed predominantly in pollens; thus, RAB2 seems to be important for pollen function in angiosperms. Dominant-negative mutants of NtRAB2 (tobacco pollen-predominant RAB2) have been shown to slow pollen-tube growth (Cheung et al., 2002), and NtRAB2 was localized to the Golgi apparatus by both fluorescent protein-tagging and immuno-electron microscopy. In addition, dominant negative mutants of NtRAB2 caused reduction in transport between the ER and the Golgi. Thus, RAB2 proteins in plants could be important regulators of ER–Golgi trafficking, as is the case with RAB2 in mammals. In a desiccation-tolerant grass, Sporobolus stapfianus, a RAB2 homolog was induced by dehydration (O’Mahony and Oliver, 1999); this suggests that the RAB2 proteins involved in ER–Golgi trafficking may also play a role in the stress response. 3.2.1.1.3. RAB6/RABH RAB6/Ypt6p is reported to function in retrograde trafficking pathways among the Golgi stacks and from the Golgi to the ER. The molecular function of RAB6 seems to be conserved in plants, because an A. thaliana RAB6 homolog (AtRABH1b) can displace the function of yeast YPT6 (Bednarek et al., 1994). It is also reported that A. thaliana RAB6/RABH proteins can physically interact with a member of the Golgin proteins, a group of proteins that act as putative tethers on the Golgi apparatus (Latijnhouwers et al., 2007). The precise function and physiological significance of plant RAB6 remains to be determined. 3.2.1.2. SNAREs SNAREs involved in ER–Golgi trafficking can be divided into two groups according to their subcellular localization: ERlocalized and Golgi-localized. In addition, some SNAREs are localized on the trans-Golgi-network (TGN); these will be discussed in Section 3.2.2 (Secretion). 3.2.1.2.1. SNAREs on the ER The ER-localization of GFP-tagged SNAREs was recognized as a reticulated fluorescence pattern throughout the cell. Uemura et al. (2004) reported six SNAREs on the ER: one Qa-SNARE, AtSYP81/AtUFE1; three Qc-SNAREs, AtSYP71,

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AtSYP72, and AtSYP73; and two R-SNAREs, AtSec22 and AtVAMP723. There were no Qb-SNAREs mapped on the ER in this study, but AtSEC20 and AtUSE1, close relatives of yeast Sec20p and Use1p, have been suggested to localize on the ER. Among these molecules, AtSYP81 was recently demonstrated to be involved in ER–Golgi trafficking. The subcellular localization of AtSYP81 suggested that this molecule predominantly resides on putative ER import sites, consistent with the idea that AtSYP81 is a Qa-SNARE responsible for retrograde transport from the Golgi to the ER (Bubeck et al., 2008). Although the physiological functions of AtSEC20 and AtSYP81/AtUFE1 remain unknown, a possible involvement in ER–Golgi trafficking has been reported. These molecules physically interact with MAG2, a protein that contains a RINT-1/TIP20 domain (Li et al., 2006). These results suggest that molecular mechanisms involving SNAREs on the ER are conserved between plants and yeast to some extent. 3.2.1.2.2. SNAREs on the Golgi apparatus The Golgi apparatus is visualized under the fluorescent microscope as a series of dot- or disk-like structures distributed throughout cytoplasm. In a systematic analysis, nine A. thaliana SNAREs: two Qa-SNAREs, AtSYP31 and AtSYP32; four QbSNAREs, AtGOS11, AtGOS12, AtMEMB11, and AtMEMB12; two QcSNAREs, AtBS14a and AtBS14b; and one R-SNARE, AtVAMP714, were mapped on the Golgi apparatus (Uemura et al., 2004). Some of these molecules have also been demonstrated to localize on the Golgi in tobacco leaf cells (Chatre et al., 2005). In addition, two SFT1 homologs were suggested to function around the Golgi, as is the case with the yeast counterpart of SFT1. Information on the molecular function of Golgi SNAREs is limited, but it has been shown that overexpression of AtSEC22, AtMEMB11, and AtSYP31 caused relocalization of a Golgiresident protein to the ER, suggesting their involvement in trafficking between the ER and the Golgi (Bubeck et al., 2008; Chatre et al., 2005). It was also reported that overexpression of AtBS14a or AtBS14b supported the growth of the yeast sft1 mutant (Tai and Banfield, 2001), indicating that at least some of the functions of Golgi-SNAREs are highly conserved among the yeast and plant kingdoms. 3.2.2. Secretion 3.2.2.1. RAB 3.2.2.1.1. RAB8/RABE Mammalian RAB8 and its yeast counterpart, Sec4, are known to mediate post-Golgi transport to the PM. In plants, a study in A. thaliana demonstrated that the RAB8/RABE group regulates vesicular trafficking in the secretory pathway, either at or after the Golgi (Zheng et al., 2005). When a dominant-negative AtRABE1 (N128I; possibly nucleotide free form) was coexpressed with secGFP, the secretion of

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secGFP was perturbed, resulting in its accumulation in cells and its misdirection to the vacuolar transport pathway. The localization of YFPAtRABE1 on the Golgi in tobacco cells also suggests that it may function in the late secretory pathway; this notion is further supported by the observation that RABE1 acts downstream of RABD2. Interestingly, expression of some proteins in this group is induced by ethylene (Moshkov et al., 2003), although the physiological significance of this finding is not clear. 3.2.2.1.2. RAB11/RABA The RAB11 group is the largest subclade in plant RABs, comprising almost half (26 members) of the total number of RABs in A. thaliana. The RAB11 group is divided into six subgroups (RABA1 to RABA6). The remarkable expansion of this group is one of the most striking features of plant RAB GTPases. The most closely related mammalian RABs, RAB11a and RAB25, have been localized to apical recycling endosomes in polarized epithelial cells. The homologous subclass in S. cerevisiae, Ypt31/32, has been implicated in endocytic recycling and vesicular export from the late Golgi compartment to the PVC or the PM. In plants, this family is likely to function in the post-Golgi pathway. Although direct evidence is sporadic on the function and localization of the RAB11 group, recent studies have shed new light on this important group. RABA1, consisting of nine members, is the largest subgroup in the RAB11/RABA group of A. thaliana. Interestingly, a close homolog has not been found in yeast or mammals. The localization and function of RABA1 have not been clearly determined in A. thaliana. However, a RABA1 protein in N. tabacum, NtRAB11b, has been shown to play a crucial role in pollen tube growth (de Graaf et al., 2005). NtRAB11b localizes to the apical clear zone of elongating pollen tubes, and the proper function of NtRAB11b is essential for secretion and endocytic recycling at the tips of pollen tubes. The RABA2 subgroup shows high similarity to mammalian RAB11 and yeast Ypt31/32, suggesting it may function in post-Golgi or endocytic pathways. On the other hand, RABA3, A4, A5, and A6 appear to have no clear counterparts in yeast or mammals. A recent study demonstrated that A. thaliana RABA2 and RABA3 colocalized on a novel post-Golgi membrane domain in root tip cells (Chow et al., 2008). The RABA2/A3 compartment was distinct from, but often close to, Golgi stacks and the PVC, and partly overlapped with the VHA-a1-positive TGN. Intriguingly, RABA2/A3 localized on cell plates that formed in dividing cells; this implies that these subgroups might play crucial roles in cytokinesis via the regulation of polarized secretion. The RABA2/A3 compartments were stained by an endocytic tracer, FM4–64, before the dye reached RAB5/RABF-positive or GNOM-positive endosomes. This result suggested that the plant TGN also bears early endosomal nature (Chow et al., 2008).

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Some lines of evidence suggest that the RABA4 subgroup in A. thaliana functions in polarized secretion in root hair cells. Enhanced yellow fluorescent protein (EYFP)-tagged RABA4b specifically localized in the tips of growing root hair cells, but was abolished in mature root hair cells that had stopped expansion (Preuss et al., 2004). The tip-localized fluorescence was disrupted by latrunculin B treatment, indicating that the proper localization of RABA4b required actin polymerization. Interestingly, the majority of this protein was found in a unique membrane compartment that did not cofractionate with the TGN SNAREs, SYP41 or SYP51 (Preuss et al., 2004). Nevertheless, RABA4b was detected on the TGN by immuneelectron microscopy (Preuss et al., 2006). These results suggest that there are at least two distinct TGN-related compartments with different protein contents. Alternatively, the TGN is a complex organelle with multiple membrane domains that are associated with different sets of proteins, and these domains migrate in different fractions in the biochemical fractionation procedure. Subcellular localization was also reported for RABA3 and RABA4 proteins in P. sativum. When expressed in tobacco BY-2 cells, GFP-tagged PRA2 (PsRABA3) and PRA3 (PsRABA4) localized on distinct populations of punctate structures that most likely included the Golgi, TGN, and PVC/endosomes (Inaba et al., 2002). There are quite a few reports on the functions of RABA5 and RABA6 proteins. Immuno-electron microscopy on pollen indicated that one member of the A. thaliana RABA5 group, ARA4 (RABA5c), localized on the Golgi and adjacent vesicular structures (Ueda et al., 1996a). The ARA4 was also used successfully in a screen for interacting molecules using the yeast system (Ueda et al., 1996b); however, the physiological function of RABA5/A6 remains to be elucidated. As mentioned above, plant RAB11 proteins reside on post-Golgi organelles including the TGN, and most likely mediate both exocytic and endocytic trafficking. This is especially important in polarized cells for trafficking events, including cell plate formation during cytokinesis and the tip growth of pollen tubes or root hairs. 3.2.2.2. SNARE 3.2.2.2.1. SNAREs on the plasma membrane (PM) In a systematic analysis, 18 A. thaliana SNAREs were localized on the PM, including: AtSYP111/KNOLLE, AtSYP112, AtSYP121 (AtSYR1, PEN1), AtSYP122, AtSYP123, AtSYP124, AtSYP125, AtSYP131, AtSYP132 (Qa), AtNPSN11, AtNPSN12, AtNPSN13, AtVTI12 (Qb), AtVAMP721, AtVAMP722, AtVAMP724, AtVAMP725, and AtVAMP726 (Qc) (Uemura et al., 2004). GFP-tag studies showed that these proteins localized mainly on the PM, and occasionally on cytoplasmic dotty organelles. Some of these punctate structures were confirmed to be endocytic compartments (for AtSYP111 and AtVAMP721) by costaining with FM4–64; others were

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confirmed to be TGN (for AtVTI12) by colocalization with other TGN SNAREs. These results suggests that SNAREs localized on the PM may be involved in both secretory and endocytic pathways. In fact, AtSYP111/ KNOLLE was suggested to be involved in both exocytic and endocytic membrane fusions; this will be discussed in more detail later. Recently, AtSYP7 members (SYP71, SYP72, and SYP73) were shown to reside on the PM (Suwastika, 2008). Proteomic analysis also identified AtSYP71 in the PM fraction (Alexandersson, 2004). These Qc-SNAREs were also found on the ER membrane. The dual localization of these molecules might imply the existence of direct trafficking route from the ER to the PM, which should be verified in future studies. Mammalian SNAP25 is known to mediate the exocytosis of synaptic vesicles (Hong, 2005). On the other hand, a transient expression analysis in protoplasts showed that GFP-tagged A. thaliana SNAP25 proteins (SNAP29, SNAP30, SNAP33) failed to localize on the PM. This was probably due to a deficiency in the posttranslational modification required for SNAP25 membrane binding. However, several lines of evidence have suggested that this subgroup functions in secretion in plants. The AtSNAP33 protein was shown to interact with KNOLLE/SYP111 (Heese et al., 2001), indicating that it may play a role in cell plate formation. AtSNAP33 was also reported to bind to AtSYP121 and its counterpart in N. tabacum, NtSyr1 (Kargul et al., 2001; Kwon et al., 2008; Leyman et al., 2000). AtSYP121 was also identified as a gene responsible for the pen1 mutation (see Chapter 4). The expression of AtSNAP33 increased upon pathogen infection and mechanical stimulation (Wick et al., 2003), suggesting it plays a role in secretion. Several other SNAREs on the PM have also been shown to be involved in the plant–microbe interaction, which will be summarized in Chapter 4. 3.2.2.2.2. Trafficking around the TGN AtSYP41/TLG2a and AtSYP42/TLG2b were the first SNAREs to be localized to the TGN. Immuno-electron microscopy further demonstrated that SYP41 and SYP42 were localized on distinct TGN domains (Bassham et al., 2000; Sanderfoot et al., 2001). These Qa-SNAREs are likely activated by AtVPS45, a Sec1prelated protein, because both AtSYP41 and AtSYP42 form complexes with AtVPS45. A systematic analysis also indicated that another Qa-SYP4 protein (AtSYP43), three Qb-VTI1 proteins (AtVTI11, AtVTI12, and AtVTI13), and Qc-AtSYP61 were also localized on the TGN (Uemura et al., 2004). These molecules form several SNARE complexes with different functions, indicating that multiple SNARE complexes execute membrane fusion at the TGN. Curiously, there have been no R-SNAREs mapped to the TGN in plant cells. AtYKT6 should be a candidate, because it mediates liposomal fusion with either AtSYP41 or AtSYP61 in vitro (Chen, 2005). However, further in vivo investigation is apparently needed.

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It is noteworthy that GFP-AtSYP41 and Venus-AtSYP61 are not always associated with the Golgi apparatus (Foresti and Denecke, 2008; Uemura et al., 2004). This raises some interesting possibilities: either not all SYP4s are localized on the TGN or some population of TGN exists independently of the Golgi apparatus. Recent studies indicate that the TGN also has early endosomal properties; this was indicated by the observation that SYP41 compartments were accessible to FM4–64 during a short period of incubation (Dettmer et al., 2006; Lam et al., 2007). These results suggest that the TGN is located at the crossroads of secretory and endocytic pathways. This idea was supported by the finding that chemical treatment with endocydin 1 caused the agglomeration of TGN residents together with some cargos bound for endocytosis, including PIN2 and AUX1 (Robert et al., 2008). However, other cargos bound for endocytosis were not affected by the chemical, suggesting that plants elaborate complex pathways of endocytosis. Further study will unveil the TGN’s function in endocytosis, the molecular machinery involved, and its physiological significance. 3.2.3. Vacuolar and PVC /endosomal trafficking 3.2.3.1. RAB 3.2.3.1.1. RAB7/RABG In mammals and yeast, RAB7 and Ypt7p are known to function in lysosomal and vacuolar biogenesis, respectively. Also in A. thaliana, most of the RAB7-related proteins, including AtRABG3b/AtRAB75, are localized on the vacuolar membrane (Saito et al., 2002; Ueda et al., unpublished results). Proteomic analysis of vegetative vacuoles from A. thaliana identified two members of this group, AtRABG3b and AtRABG3d/AtRAB72 (Carter et al., 2004). Interestingly, the same proteomic study also identified other RAB group members in the vacuolar membrane fraction, including RABD2a/ARA5, RABE1c/ ARA3, RABB1a, and RABD2b. Some of these molecules are proposed to function in ER–Golgi trafficking, which may reflect a direct trafficking route from the ER to vacuoles in plants (Hara-Nishimura et al., 1998). Although the physiological significance of the RAB7 group is not yet crystal clear, it is reported that the overexpression of AtRABG3e in plants confers tolerance to salt and osmotic stress (Mazel et al., 2004). This result suggests that the RAB7/RABG group plays important roles in the adaptation to abiotic stress through the regulation of vacuolar functions. 3.2.3.1.2. RAB5/RABF RAB5 is the best-characterized RAB GTPase in mammalian cells. Mammalian RAB5 localizes on the PM and early endosomes, and regulates a wide spectrum of endocytic events, including the biogenesis of clathrin coated vesicles, homotypic fusion between early endosomes, endosomal motility, compartmentalization of endosomal membrane domains, modification of lipid moieties in endosomal membranes, and endosomal signaling (Bucci et al., 1992; Christoforidis

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et al., 1999; Gorvel et al., 1991; Hoepfner et al., 2005; Miaczynska and Zerial, 2002; Miaczynska et al., 2004; Nielsen et al., 1999; Shin et al., 2005). RAB5 is widely conserved in plants; all green plant lineages whose genomes were fully sequenced contained orthologous genes to the mammalian RAB5. In addition to these RAB5 orthologs, plants harbor a plantunique type of RAB5 that has several outstanding features in its structure. This unique group of RAB5 proteins lacks a C-terminal hypervariable region and the adjoining Cys motif, which are essential for localization, membrane binding, interaction with GDI, and the function of conventional RAB GTPases. Instead, they harbor an extra amino acid stretch at the N-terminus that can be fatty acylated in order to anchor the protein to the membrane. The first plant-unique RAB5 was identified in L. japonicus (Borg et al., 1997), followed by its identification in other plants, including Mesenbryanthemum crystallinum (Bolte et al., 2000) and A. thaliana (Ueda et al., 2001). One of the most prominent features in the overall organization of plant RAB GTPases is that two RAB5 groups are conserved among plants. There are three RAB5 homologs in the A. thaliana genome, including ARA7/RABF2b, RHA1/RABF2a, and ARA6/RABF1. ARA7 and RHA1 are conventional RAB5s, and ARA6 is a plant-unique RAB5. All these RAB5 proteins are in the endocytic pathway, as organelles bearing these RAB5s could be stained with FM4–64 (Ueda et al., 2001, 2004), a robust marker for endocytic compartments in plant cells (Griffing, 2008; van Gisbergen et al., 2008). The endocytic nature of the ARA7-compartments was also confirmed in a study using BOR1, a boron transporter whose endocytosis is induced by a high supply of boron (Takano, 2005). During endocytosis, BOR1 passed through ARA7-endosomes before reaching the vacuoles. In addition, functional analysis with a dominant negative ARA7 indicated an involvement in endocytosis; when the dominant negative ARA7 was overexpressed in A. thaliana plants and tobacco BY-2 cells, endocytosis of FM4–64 was severely affected (Dhonukshe et al., 2006). Furthermore, several lines of evidence have suggested that plant RAB5s also function in the vacuolar transport pathway. The overexpression of dominant negative mutants of ARA7 and RHA1 disturbed the transport of soluble vacuolar proteins to the vacuole in N. tabacum leaf cells and A. thaliana protoplasts, respectively (Kotzer et al., 2004; Sohn et al., 2003). These results indicate that conventional RAB5s come into play at the point where endocytic and biosynthetic pathways are closely associated or merged. Recently, ultrastructural analysis by immuno-electron microscopy demonstrated that all RAB5s localized on multivesicular bodies (Haas et al., 2007), suggesting that at least some portion of the RAB5-bearing endosomes corresponded to the organelle previously identified as the prevacuolar compartment. This evidence is also consistent with the observation that RAB5 proteins mediate both endocytic and vacuolar transport.

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Contradictory results have been reported on the participation of plantunique RAB5 proteins in vacuolar transport. The overexpression of dominant negative ARA6 did not perturb the transport of sporamin-GFP to the vacuole (Sohn et al., 2003). On the other hand, the dominant negative construct of m-RABmc, a close homolog of ARA6 in M. crystallinum, conferred an inhibitory effect on the trafficking of aleurain-GFP to the vacuole (Bolte et al., 2000). This discrepancy might be explained by the fact that sporamin is a storage protein in tubers, while aleurain-GFP is sorted to the lytic vacuole. Nevertheless, all of the results above were obtained from transient expression experiments; thus, to clarify the precise functions of these molecules, further studies should be conducted under more physiological conditions and genetic analyses should be performed using loss-offunction mutants. 3.2.3.1.3. RAB18/RABC It has been suggested that the mammalian RAB18 protein is implicated in multiple cellular functions, including endocytic transport (Lutcke et al., 1994), lipid droplet localization (Ozeki et al., 2005), and secretion (Vazquez-Martinez et al., 2007); however, its precise function is not yet clear. There have been no reports on the localization or function of the plant RAB18 proteins. Because the amino acid sequences of plant and animal RAB18 proteins differ substantially in the conserved domains that define subclass specificity, it is possible that the plant RAB18 proteins might have totally different functions from animal RAB18 proteins. This remains to be clarified by future studies. 3.2.3.2. SNARE A systematic analysis revealed that nine SNARE molecules localized on the vacuolar membrane, including: Qa-AtSYP21, Qa-AtSYP22, Qb-AtVti11, Qb-AtVti13, Qc-SYP51, Qc-SYP52, R-VAMP711, R-VAMP712, and R-VAMP713 (Uemura et al., 2004). AtSYP21 (AtPEP12) was the first SNARE to be isolated from plants (Bassham et al., 1995), and its PVC localization was later demonstrated by immuno-electron microscopy (da Silva Conceicao et al., 1997). In protoplasts, AtSYP21 exhibited a predominant dot-like pattern and occasionally a weak staining on vacuolar membranes (Uemura et al., 2004). In contrast, AtSYP22 mainly localized on the vacuolar membrane, and sometimes on the dot-like structures that are apparently associated with the vacuolar membrane. Thus, two homologous Qa-SNAREs, AtSYP21 and AtSYP22 localized on the vacuolar membrane and the PVC with different densities. The AtSYP21 and AtSYP22 genes were isolated by complementation of different yeast mutants (pep12 and vam3, respectively) (Bassham et al., 1995; Sato et al., 1997); moreover, a functional analysis by overexpression in protoplasts indicated that these gene products had distinct functions (Foresti et al., 2006). On the other hand, a molecular phylogenetic analysis suggested that these genes were the product of a gene duplication that

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occurred independently of the gene duplication in yeast that yielded PEP12 and VAM3 (Dacks et al., 2008). It would be interesting to examine the functional differences of AtSYP21 and AtSYP22 in a more physiological experimental system. AtSYP23 is a homologous Qa-SNARE expressed in A. thaliana. AtSYP23 is functionally redundant with AtSYP22 (Ohtomo et al., 2005), but is not fully functional due to a deleterious polymorphism in some A. thaliana accessions, including Columbia (Zheng et al., 1999a). The fluorescence patterns of GFP-tagged AtVTI11, AtVTI13, AtSYP51, AtSYP52, AtVAMP711, AtVAMP712, and AtVAMP713 were similar to that of GFPAtSYP22. The vacuolar localizations of SYP51, SYP52, Vti11, and SYP22 were also supported by a proteomic analysis of vacuolar membranes (Carter et al., 2004). To date, AtVAMP727 is the only SNARE known to predominantly localize on the endosomes/PVC. The organelles bearing AtVAMP727 were stained by FM4–64 (Uemura et al., 2004), indicating that AtVAMP727 is on endocytic organelles. The colocalization of AtVMAMP727 with RAB5 proteins (ARA7 and RHA1) also suggests its endosomal localization (Ueda et al., 2004). Recently, the four components that comprise the AtVAMP727 complex were identified as: AtSYP22, AtVTI11, AtSYP51, and AtVAMP727 (Ebine et al., 2008). The AtVAMP727 gene exhibited intimate genetic interactions with the AtSYP22 gene, including synthetic lethality and multicopy suppression. Furthermore, the protein products of these genes colocalized at the PVC subdomain that seems to be associated with the vacuole. These results strongly suggest that this SNARE complex executes membrane fusion between PVCs and vacuoles; however, VAMP727 may also be involved in other fusion events because some VAMP727 was found on VAM3-free PVCs. 3.2.4. Cytokinesis 3.2.4.1. RAB It was not clear which RABs are involved in membrane targeting and fusion on newly forming cell plates. Recently, however, RAB11/RABA proteins were reported to play a role. RABA2 and RABA3 were found on a novel post-Golgi membrane domain (Chow et al., 2008) and in the mitotic phase they colocalized with KNOLLE/ AtSYP111 on cell plates. In particular, these proteins localized to the growing edges of the cell plates, where VHA-a1, GNOM, and PVC residents were excluded. Conditional expression of a dominant-negative RABA2a construct resulted in enlarged polynucleate cells with cell wall stubs. This indicated that RABA2/A3 must play a critical role in cell plate formation; presumably, it regulates secretion or endocytosis associated with developing cell plates.

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3.2.4.2. SNARE KNOLLE AtSYP111 was identified as the gene responsible for the knolle mutant, which is defective in the establishment of polarity in the embryo (Lukowitz et al., 1996; Mayer et al., 1991). KNOLLE was proposed to regulate membrane fusion during cytokinesis based on the facts that KNOLLE is expressed in a cell cycle-dependent manner and knolle mutants frequently exhibit multinucleation and incomplete cell wall formation (Lukowitz et al., 1996). In addition, KNOLLE was shown to localize on forming cell plates, and the knolle mutant exhibited many unfused vesicles at the predicted sites of cell plate formation (Lauber et al., 1997). These results suggested that KNOLLE is a Qa-SNARE that specifically executes cytokinetic vesicle fusion. Yeast two-hybrid screening was performed to identify proteins that interacted with KNOLLE. AtSNAP33 was identified as a partner of KNOLLE (Heese et al., 2001). AtSNAP33 colocalized with KNOLLE on cell plates, and an AtSNAP33 loss of function resulted in incomplete cytokinesis. Two other SNAP25-like proteins (AtSNAP29 and AtSNAP30) also interacted with KNOLLE in the yeast two-hybrid assay, suggesting these molecules may also function in cytokinesis. NPSN11 is another SNARE that is likely to function cooperatively with KNOLLE in cytokinesis (Zheng et al., 2002). NPSN11 cofractionated with KNOLLE in fractionation analysis, and immunofluorescence microscopy showed that it was localized to cell plates in dividing cells. It has been proposed that cell plates are formed by the fusion of newly forming cell plates with Golgi-derived vesicles that harbor membrane and cell wall components. However, a recent proposal suggests that endocytic vesicles contribute to cell plate formation (Dhonukshe et al., 2006). It has been shown that endocytosis was strongly enhanced during cell plate formation; moreover, cell surface materials were rapidly delivered to forming cell plates, including PM proteins, cell wall components, and exogenously applied endocytic tracers. Currently, researchers in the field have not fully agreed on the significance of the endocytic pathway in cytokinesis (Reichardt et al., 2007). The answer to this question awaits further analysis. We suggest that new methodologies, including specific inhibition of endocytosis in conditional mutants or the use of specific inhibitors would be good approaches for clarifying the role of endocytosis in cytokinesis.

4. RABs and SNAREs in Higher Order Plant Functions In the previous chapter, we discuss how RABs and SNAREs play critical roles in various plant functions, from basic ER–Golgi trafficking to plant-specific cytokinesis. Recent studies have also revealed that many other plant functions involve RABs and SNAREs. Forward genetic techniques

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Cell differentiation, Flowering, Morphogenesis SYP22

SYP23 Gravitropism SYP22

VTI11 Abiotic stress SYP61

VTI12

VAMP711

RAB7 RABG

Tip growth Pollen tube RAB2 RABB

Plant-microbe interaction Immune system SYP121

VAMP721

SYP122

VAMP722

SYP32

SNAP34

RAB11 RABA

Root hair RAB11 RABA

Autophagy VTI12

RAB5 RABF

Symbiosis RAB1 RABD

RAB7 RABG

Figure 4.12 RABs and SNAREs are involved in various important higher order plant functions. Here, the RABs and SNAREs of A. thaliana are categorized according to their involvement with a particular plant function.

with A. thaliana have accelerated investigations into the roles these molecules play in higher order plant functions. In addition, current reverse genetic techniques provide essential tools for attaining an in-depth understanding of the mechanisms of action for these proteins. In this chapter, we will summarize recent progress in studies on the functions of RABs and SNAREs in complex and higher order functions (Fig. 4.12).

4.1. Gravitropism The direction of plant growth turns in response to gravity; this is called gravitropism. Stems show negative gravitropism, thus the arial parts of plants grow upward. To unveil the molecular mechanisms involved in shoot

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gravitropism, A. thaliana mutants (shoot gravitropism: sgr) that are defective in the shoot response to gravity have been screened (Fukaki et al., 1996; Yamauchi et al., 1997). The sgr1 gene was found to be an allele of a gene that encodes the GRAS transcription factor, SCARECRAW (SCR). Impairments in SCR resulted in loss of the endodermal layer responsible for sensing gravity (Fukaki et al., 1998). Subsequently, the genes responsible for zig/sgr4 and sgr3 were identified, and the results were surprising for researchers in the vesicular trafficking field. ZIG/SGR4 and SGR3 encode Qb-AtVTI11 (Kato et al., 2002) and Qa-AtSYP22/VAM3 (Yano et al., 2003), respectively. Both gene products localize on vacuoles, and vacuolar morphology is disorganized in endodermal cells in zig/sgr4 and sgr3 mutants. These mutants showed abnormal amyloplast mobility in endodermal cells, which are known to act as statocysts for gravity sensing (Saito et al., 2005). The mutant phenotypes were restored to normal when these genes were expressed under the regulation of an endodermis-specific SCR promoter (Morita et al., 2002; Yano et al., 2003). These results indicate that the vacuolar function in endodermis is critical in an early step of the gravitropic response. Suppressor mutants of zig-1 (zig suppressor; zip) were then screened to investigate the molecular basis of ZIG/AtVTI11-mediated vesicular transport and gravitropism. In addition to the defect in gravitropism, zig/sgr4 showed morphological abnormalities, including zigzag-shaped inflorescences and wavy leaves. A dominant mutant, zip1, rescued both the gravitropic and morphological defects of zig-1 (Niihama et al., 2005). The zip1 mutation was caused by single amino acid substitution in AtVTI12, a Qb-SNARE closely related to AtVTI11. This amino acid substitution affected the localization of AtVTI12, which allowed an ectopic SNARE complex formation to take over the function of the defective AtVTI11. Although it is predicted that some RAB GTPases cooperate with vacuolar SNAREs to maintain vacuolar membrane integrity, no RABs have been shown to be involved in gravitropism. This might be due to the functional redundancy in RAB subfamilies, although it is possible that SNAREs might play some unique roles independently of RAB GTPases in the gravitropic response.

4.2. Tip growth Pollen tubes and root hairs show characteristic polar cell expansion called tip growth. An underlying mechanism for rapid, focused tip growth is thought to be efficient, regulated membrane trafficking that incorporates membrane and deposits cell wall components at the tube apex. Recent studies have revealed that RAB is an essential spatio-temporal regulator of vesicular trafficking in growing tips.

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4.2.1. Pollen tube growth Tip-growth occurs when pollen tubes elongate within the female reproductive tissues to deliver sperm cells to ovules for fertilization. Two tobacco RAB GTPases have been reported to play critical roles in this process, NtRAB2 and NtRAB11b. NtRAB2, a Golgi-localized RAB2/RABB protein, is expressed predominantly in pollens (Cheung et al., 2002). Expression of a dominant-negative form of NtRAB2 perturbed ER– Golgi trafficking in pollen tubes and inhibited the proper transport of cargo proteins and tube growth. Interestingly, NtRAB2 was not properly recruited to the Golgi apparatus when expressed in leaf cells. This result suggested that the pollen tube may possess a specialized mechanism for localizing NtRAB2 to the Golgi. NtRAB11b is the other RAB GTPase demonstrated to play a role in pollen tube growth (de Graaf et al., 2005). NtRAB11b predominantly localizes on vesicles concentrated at the apical clear zone of elongating pollen tubes. These vesicles require an intact actin cytoskeleton for correct localization. A functional GTPase cycle also seems to be essential for the correct localization of vesicles in pollen tubes. The efficient accumulation of NtRAB11b-positive vesicles in the apical clear zone was abolished with the expression of either a dominant-negative (DN; presumably GDP-fixed form) or a constitutively active (CA; GTP-fixed form) NtRAB11b. Furthermore, overexpression of either DN or CA NtRAB11b inhibited pollen tube growth both in vitro and in vivo, and consequently reduced fertility. 4.2.2. Root hair growth Another tip growth event that involves RAB11/RABA is the growth of root hair. AtRABA4b is ubiquitously expressed in plant tissues, including roots, leaves, stems, and flowers. When AtRABA4b was tagged with GFP and expressed under the control of a constitutive 35S promoter, it localized to novel organelles at the tips of growing root hair cells in an actindependent fashion. Tip localization of AtRABA4b-domains also required the expression of some root hair defective (RHD) gene products, indicated by their mislocalization in rhd mutants. Recently, one of the mutated rhd genes, RHD4, was found to encode a phosphatidylinositol-4-phosphate [PI (4)P] phosphatase. PI(4)P localizes in the PM at the tip region of growing root hair; this suggests that PI(4)P, or one of its derivatives (e.g., PI(4,5)P2, generated by phosphatidylinositol-phosphate 5-kinase), may act as a positional cue at the tip region (Kusano et al., 2008; Thole et al., 2008). This finding gave rise to the question of whether there was a direct linkage between phosphoinositide metabolism and RAB11/RABA functions. A study on the AtRABA4b effector molecule provided an important insight into this question.

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A yeast two-hybrid screen revealed that phosphatidylinositol 4-OH kinase (PI4Kb1) interacted with GTP-bound AtRABA4b (Preuss et al., 2006). PI4Kb1 and AtRABA4b colocalized at the tip region of growing root hair. Additionally, a double mutant that reduced the function of PI4Kb1 and its close relative, PI4Kb2, exhibited aberrant root hair development. These lines of evidence suggested that AtRABA4b regulates the activity or localization of PI4Kb, or vice versa, and through this interaction, tip localization of phosphoinositides is established or maintained. There is currently no experimental evidence indicating the function of SNAREs in tip growth. However, microarray data available in the public database indicates that some SNAREs, including AtSYP124, AtSYP125, and AtSYP131 are expressed exclusively in pollens. This observation suggests that active and specialized SNARE-mediated trafficking may play a crucial role in pollen tube growth, but this hypothesis awaits confirmation with future functional studies.

4.3. Autophagy Several lines of evidence have suggested a link between vacuolar/endosomal trafficking and autophagy. When grown under normal conditions, atvti12 mutant plants do not show any abnormal phenotypes, but atvti11/ zig exhibits easily recognizable phenotypes, including zigzag-shaped inflorescence and wavy leaves. AtVTI11 and AtVTI12 seem to share redundant functions; it has been shown that the double mutant is lethal and the elevated expression of AtVTI12 suppressed a zig-1 mutation. On the other hand, functional analysis using a yeast vti1 mutant demonstrated that only AtVTI12 had the ability to restore a defect in the cytoplasm-tovacuole (Cvt) pathway. This suggested that VTI1 proteins have distinct functions, which was recently confirmed by functional analyses in plants (Sanmartin, 2007). In the detached-leaf assay, atvti12 showed accelerated leaf senescence, a phenotype typical for the autophagy-defective mutants, atapg7 and atapg9 (Hanaoka et al., 2002). Electron microscopy also indicated that an autophagy-like process was affected in the cells of atvti12. Accumulation of authophagosome-like structures was also observed in the embryos of A. thaliana vcl1 and vps9a-1 mutants that have impairments, respectively, in the VPS16 homolog and in the gene encoding a GEF for RAB5/RABF proteins (Goh et al., 2007; Rojo et al., 2001). Both of these mutants exhibited embryonic lethality; but otherwise, they had distinct phenotypes. This suggested that VCL1 and VPS9a act in distinct trafficking pathways. Thus, these gene products might not play a direct role in autophagy, but it is possible that vacuolar/endosomal trafficking is indispensable for the normal progression of autophagy in plants.

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4.4. Plant–microbe interaction 4.4.1. Plant immune system Pathogenic fungi can breach the plant cell wall when they are settled onto the cell surface, but they cannot penetrate into ‘‘nonhost’’ plant cells. In the course of a molecular genetics study regarding nonhost resistance, a SNARE was identified as a key component in this system (Collins et al., 2003). The penetration ( pen) mutants of A. thaliana are defective in their resistance against the nonhost penetration of barley powdery mildew, Blumeria graminis f. sp. hordei. Among pen mutants, one of the mutated genes, PEN1 was demonstrated to encode a Qa-SNARE, AtSYP121/ AtSYR1. The AtSYP121 gene is an A. thaliana homolog of the ROR2 gene in barley (Hordeum vulgare), which is also required for basal penetration resistance against B. g. f.sp.hordei. Both AtSYP121 and ROR2 are localized on the PM in uninfected cells, and upon inoculation, they both relocate and accumulate at penetration sites. Recently, other components of the SNARE complex were identified, and it was found that this SNARE complex was comprised of AtSYP121, AtSNAP33 (a protein encoded by the homolog of HvSNAP34 that is also required for H. vulgare immunity), and either AtVAMP721 or AtVAMP722 (Kwon et al., 2008). HvSNAP34 was also shown to form a complex with ROR2. AtSYP122 is a close relative of AtSYP121, which is also responsive to infection (Assaad et al., 2004). Genetic analysis indicated that these two SYP12 proteins have different but partially overlapping activities. While the PEN1/AtSYP121 protein is mainly responsible for secretion during papilla formation, the AtSYP122 protein seems to function in various secretion pathways, including cell wall deposition (Assaad et al., 2004). Subsequent studies further demonstrated that these proteins negatively regulated the downstream defense pathway mediated by salicylic acid, jasmonate acid, and ethylene signaling pathways (Zhang et al., 2007b). 4.4.2. Symbiosis and nodule formation It was suggested 15 years ago that vesicular trafficking probably contributes to symbiosis and nodule formation in legumes. The peribacteroid membrane (PBM) surrounding rhizobium in nodules originates from the fusion and expansion of newly synthesized vesicles with the PM. To explore the function of RAB GTPases in PBM biogenesis and nodule development, several genes encoding RAB GTPases were isolated from legumes (sRAB1 and sRAB7 from G. max and vRAB7 from V. aconitifolia) (Cheon et al., 1993). Elevations were observed in sRAB1 and vRAB7 gene expression during nodule formation, and antisense suppression of these genes affected different steps of nodule development. Antisense suppression of sRAB1 resulted in a reduced number of bacteroids in the cytoplasm and vacuoles; antisense suppression of vRAB7 resulted in the accumulation of late

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endosomal structures, including multivesicular bodies in the perinuclear region. Recently, two Sed5p-related Qa-SNAREs, LjSYP32–1 and LjSYP32–2 were isolated in L. japonicus. The antisense suppression of these genes also causes abnormal nodule development and retardation of plant growth (Mai et al., 2006). Together, these data indicate that RABs and SNAREs play critical roles in symbiosis and nodule formation, although the precise mechanisms involved remain unclear.

4.5. Abiotic stress response A few SNARE molecules have been shown to be involved in salinity and osmotic stress tolerance. Forward molecular genetic studies found that a mutation in the AtSYP61 gene was involved in the phenotype of the osmotic stress-sensitive mutant (osm1). The osm1 exhibits increased sensitivity to both ionic (NaCl) and nonionic (mannitol) osmotic stress. osm1 plants are more apt to wilt than wild types when grown in soil with limited moisture; this is probably due to the impaired ABA-induced stomata closure. On the other hand, salt stress tolerance was conferred by the antisense suppression of the vacuolar R-SNARE, AtVAMP711 (Leshem et al., 2006). A deficiency in AtVAMP711 caused inhibition of the fusion of H2O2containing vesicles and a subsequent accumulation of H2O2-containing megavesicles in the cytoplasm. This resulted in increased salt tolerance, probably mediated by calcium-dependent protein kinases. Although the precise roles of SNAREs in stress responses remain to be elucidated, these results clearly indicate that SNARE-mediated vesicle fusion plays an important role in multiple stress responses.

4.6. Cell differentiation, morphogenesis, and flowering As mentioned above, a single amino acid substitution in AtSYP22/VAM3/ SGR3 caused the loss of shoot responsiveness to gravity (Yano et al., 2003). Surprisingly, other mutations in AtSYP22 have totally different effects. A peptide insertion in the AtSYP22 protein expressed in the short stem and midrib (ssm) mutant, caused semi-dwarfism and wavy leaf morphology due to reduced and/or unregulated cell elongation/expansion (Ohtomo et al., 2005). The ssm mutation can be suppressed by the Landsberg erecta (Ler) allele of AtSYP23, a close homolog of AtSYP22, but not by the Columbia allele of AtSYP23, which has an innate mutation that impairs its function. Interestingly, atsyp22/vam3 mutants also exhibit excessive idioblast differentiation associated with the hyperaccumulation of a myrosinase encoded by TGG1 and TGG2 (Ueda et al., 2006). This result suggests that AtSYP22/VAM3 is involved in the determination of cell fate. Furthermore, atsyp22/vam3 mutants exhibit a late flowering phenotype apparently caused by a defect in the autonomous flowering promotion pathway (Ebine and

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Ueda, unpublished results). Further studies are needed to investigate how this SNARE is implicated in the diverse phenomena of cell expansion, cell fate determination, and flowering.

5. A Unique Regulatory System in Plants for Endocytic/Vacuolar Transport Up to this point, we have mapped RAB and SNARE proteins to their sites of action in the trafficking network, summarized the links between those proteins and plant functions, and outlined how plants have recruited RABs and SNAREs for a plant-unique way of life. In the last part of this review, we would like to introduce the plant-unique ‘‘machinery’’ that has evolved for the regulation of endocytic/vacuolar transport pathways, with a special focus on RAB5. In A. thaliana, ARA7/RABF2b and RAH1/RABF2a are orthologs of the mammalian RAB5 gene, while ARA6/RABF1 encodes a plant-unique type of RAB5 with several unique features. ARA6 lacks the usual C-terminal Cys motif followed by the hypervariable region known to be essential for interactions with GDI, correct subcellular localization, and other functions. Instead, ARA6 harbors an extra amino acid stretch at the N-terminus that provides a site for N-myristoylation and palmitoylation. This dual fatty acylation is essential for its localization to endosomes; a mutation in either of the fatty acylation sites causes mislocalization of this protein. ARA6 homologs are found only in land plants, which might imply that they are involved in land plant-specific functions. ARA6 and the two other RAB5 proteins seem to localize on different, but overlapping, populations of endosomes/PVCs. In protoplasts that carry the gnom mutation, ARA6-positive endosomes have unaffected morphology, but ARA7- and RHA1-positive endosomes are deformed (Geldner et al., 2003; Ueda et al., 2004). These results suggest that ARA6 and conventional RAB5s perform distinct functions. Despite the expected differences in function, these three RAB5 proteins are activated by a single type of GEF, AtVPS9a; the VPS9 domain contains a catalytic core that is conserved in RAB5 GEFs (Goh et al., 2007). A loss of function in AtVPS9a causes embryonic lethality. Phenotypic analysis of atvps9a mutants indicated that AtVPS9a is involved in various cellular activities, including cytokinesis, paramural body formation, autophagy, and establishment of cell polarity. Furthermore, leaky atvps9a-2 phenotypes indicate that AtVPS9a plays critical roles in plant development and root growth. Overexpression of constitutively active ARA7 suppresses abnormal phenotypes of atvps9a-2; this supports the notion that AtVPS9a activates ARA7 in vivo. In contrast, the overexpression of ARA6 with an equivalent

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mutation does not suppress abnormal phenotypes of atvps9a-2. This result strongly suggests that plant-unique ARA6 functions differently than conventional ARA7 and RHA1 proteins. In recent studies, we found that the function of ARA6 counteracts that of conventional RAB5s in multiple steps of plant development (our unpublished results). It would be quite important and interesting to investigate why and how land plants have evolved this unique RAB5-based regulation of endocytic/vacuolar transport pathways. In mammalian cells, RAB5s are activated at multiple steps in the endocytic pathway by distinct GEF proteins, including Rabex-5, RIN, Rme-6, ALS2, and ANKRD27. In contrast, in A. thaliana cells, RAB5s are activated by practically a sole GEF, AtVPS9a (one other RAB5 GEF is found in A. thaliana, AtVPS9b, but its expression is limited to gametes; our unpublished results). Thus, plants seem to have employed a mechanism for activating RAB5 that is distinct from other organisms. It should also be noted that plants do not harbor genes orthologous to the mammalian RAB4 or RAB9 genes, which also regulate the endocytic pathway in mammalian cells. These lines of evidence indicate that plants have evolved unique endocytic machineries. In addition, RAB5 proteins also play critical roles in the vacuolar transport pathway (Bolte et al., 2000; Kotzer et al., 2004; Sohn et al., 2003). Because great complexity is observed in the plant vacuolar system (Bassham and Blatt, 2008; Becker, 2007; Frigerio et al., 2008; Martinoia et al., 2007), plants might also require distinct vacuolar trafficking pathways involving RAB5 proteins. VAMP727 is another good example of a plant-unique molecule involved in endocytic/vacuolar transport pathways. The most unique structural property of VAMP727 is an acidic insertion consisting of 19 amino acids in its longin domain (Uemura et al., 2004) (Fig. 4.9). Recently, we have found that the complex containing VAMP727 is essential for A. thaliana seed maturation (Ebine et al., 2008). This result is consistent with the notion that the VAMP727-like R-SNARE is conserved only in seed plants, and suggests that VAMP727 might accelerate the evolution of the seed, and adaptation of plants to land life.

6. Concluding Remarks and Future Issues RAB and SNARE proteins are conserved key components involved in vesicular trafficking. They play essential roles in common eukaryote events and in plant-unique functions. In particular, plant post-Golgi trafficking pathways show remarkable complexity, suggesting a tight linkage between divergent post-Golgi functions and a plant-unique way of life. Although our understanding of plant vesicular trafficking is rapidly expanding, many questions still remain unanswered. For example, how

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did plants develop plant-unique trafficking pathways, including direct transport from the ER to the vacuole? Are these pathways realized by recruiting conserved RABs and SNAREs? How do plants modify the conserved machinery to carry out higher order plant-unique functions? And more basically, do conserved molecules function the same way in animals, yeast, and plants? In addition to RABs and SNAREs, the associated regulators and downstream effectors will be examined in more biochemical and genetic detail in the near future. Molecular genetics approaches will continue to unveil surprising links between higher order plant functions and intracellular vesicular trafficking. Comparative genomics will broaden our evolutionary perspective, and systems biology will continue to offer a powerful methodology for understanding complicated biological phenomena. Integrative analysis using a multidisciplinary approach will enable the investigation of the fascinating questions mentioned above.

ACKNOWLEDGMENTS The authors thank Tomohiro Uemura for critical reading, and Akiko Furuyama for her help in preparing the manuscript. This work was supported by Grants-in-Aid for Scientific Research and the Targeted Proteins Research Program (TPRP) from the Ministry of Education, Culture, Sports, Science and Technology of Japan, the RIKEN Bioarchitect Research project, and the RIKEN Extreme Photonics Research project.

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Rab Proteins and Their Interaction Partners Angelika Barnekow, Anika Thyrock, and Daniel Kessler Contents 236 242 242 244 246 253 257 265 265 265

1. Introduction 2. Rab Proteins 2.1. Rab family 2.2. GTPase cycle of Rab proteins 2.3. Structure and phylogeny of Rab GTPases 2.4. Rab GTPases and malignancies 3. Small GTPases Rab1, Rab6, and Their Effectors 4. Concluding Remarks Acknowledgments References

Abstract The Ras superfamily consists of over 150 low molecular weight proteins that cycle between an inactive guanosine diphosphate (GDP)-bound state and an active guanosine triphosphate (GTP)-bound state. They are involved in a variety of signal transduction pathways that regulate cell growth, intracellular trafficking, cell migration, and apoptosis. Several methods have been devised to detect and characterize the interacting partners of small GTPases with the aim of better understanding their physiological function in normal cells and tumor cells. The Rab (Ras analog in brain) proteins form the largest family within the Ras superfamily. Rab proteins regulate vesicular trafficking pathways, behaving as membrane-associated molecular switches. The guanine nucleotide-binding status of Rab proteins is modulated by three different classes of regulatory proteins, which have been extensively studied for the Rab molecules but also for other subfamilies of the Ras superfamily. Furthermore, numerous effector molecules have been isolated especially for the Rab subfamily of proteins, which interact via a Rab-binding domain (RBD) and are recruited afterwards to specific sub-cellular compartments by the Rab proteins.

Department of Experimental Tumorbiology, University of Mu¨nster, Badestraße 9, 48149 Mu¨nster, Germany International Review of Cell and Molecular Biology, Volume 274 ISSN 1937-6448, DOI: 10.1016/S1937-6448(08)02005-4

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2009 Elsevier Inc. All rights reserved.

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1. Introduction The Ras superfamily of small GTPases was named after the rat-derived Harvey sarcoma virus oncogene ras (Karnoub and Weinberg, 2008). Small GTPases act as molecular switches in a variety of cellular processes. They are related to the a-subunit of large heterotrimeric GTPases, but function as monomeric GTP-binding proteins. The GTP-binding proteins have at least one structural element in common, the so-called G-domain, binding guanine nucleotides and hydrolyzing GTP. The basis for their function is a cycling between a GTP- (‘‘switch on’’) and a guanosine diphosphate (GDP)-bound (‘‘switch off ’’) form. As the GTPase activity of the small GTPases is intrinsically very low and thus the hydrolysis from GTP to GDP (‘‘turning off the switch’’) needs to be accelerated by additional GTPase activating proteins (GAPs) (see Section 2.2), these proteins are often termed small GTP-binding proteins or simply G-proteins. The G-domain of all small GTPases is build up of six b-sheets and five a-helices (Vetter and Wittinghofer, 2001). The major structural changes between the GTP- and GDP-bound states appear within two regions termed switch I and switch II. Differences in structure and posttranslational modifications determining their subcellular localization enable the small GTPases to act as regulators in a wide range of cellular processes (Biou and Cherfils, 2004; Wennerberg et al., 2005). The Ras superfamily is divided into six families based on structural and functional aspects, namely Ras, Rho (Ras homologous), Ran (Ras-like nuclear), Sar1 (Secretion-associated and Ras-like)/Arf (ADP ribosylation factor), RGK (Rad, Gem, Kir), and Rab (Ras analog in brain) (Chavrier and Goud, 1999; Finlin et al., 2000; Karnoub and Weinberg, 2008; Pfeffer, 2007; Wennerberg et al., 2005; Table 5.1). The eponymous proteins for the whole superfamily, the members of the Ras family, are involved in signalling cascades that regulate gene expression and cell morphology as an answer to extracellular stimuli. They have been within the focus of extensive research due to their involvement in human tumorigenesis (Karnoub and Weinberg, 2008; Macara et al., 1996; Wennerberg et al., 2005). The Rho family contains around 25 human proteins, the best known of which are RhoA, Rac1, and Cdc42 (Ridley, 2004; Wennerberg and Der, 2004). Rho proteins function in a broad range of fundamental processes like organization of the actin cytoskeleton, cell morphology and movement, or gene expression (Etienne-Manneville and Hall, 2002; Wennerberg and Der, 2004). The Rho-family members Miro 1 and 2, which form a distinct subgroup within the family, play a role in the kinesin-driven transport of mitochondria along microtubules (Boldogh and Pon, 2007; Wennerberg and Der, 2004). The Ran GTPase forms its own branch consisting only of

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Table 5.1 The families of small GTPases: Their role and function GTPase families

Roles and functions

Ras family

Ras proteins activate ERK (extracellular-signal regulated) kinases and regulate gene transcription, cell growth, and cellular transformation Rho proteins activate Rho kinases and regulate cytoskeleton-linked processes The Sar/Arf proteins are involved in protein trafficking and modulate vesicle budding and uncoating within the Golgi The Ran protein binds nuclear pores and regulates nuclear import and export RGK proteins function as potent inhibitors of voltage-dependent calcium channels. Gem and Rad modulate Rho-dependent remodeling of cytoskeleton Rab proteins are involved in vesicular transport processes (endo- and exocytosis)

Rho family Sar1/Arf family

Ras-like nuclear (Ran) protein RGK (Rad, Gem, Kir) family

Rab family

this one protein, but it is the most abundant GTPase in the cell. It is involved in the regulation of macromolecular transport between the nucleus and the cytoplasm (Weis, 2003). The Arf family GTPases, including the Arl (Arf-like) and Sar proteins, participate in vesicle formation and intracellular transport events (Pasqualato et al., 2002). The RGK (Rad, Gem, Kir) proteins constitute a novel subfamily of small Ras-related GTPases that functions as potent inhibitor of voltagedependent Ca2+ channels (VDCC) and regulator of actin cytoskeletal dynamics (Correll et al., 2008). Rab proteins represent the largest subgroup within the Ras superfamily with more than 60 members, including alternatively spliced forms, identified in human cells. In Table 5.2, the majority of the Rab proteins known so far is summarized, their swiss-prot entry number is given and some manifestations on their cellular localization and involvement in cellular transport processes are presented. Rab proteins are the key regulators in endo- and exocytotic transport processes and control membrane traffic as well as the identity of cellular compartments (Zerial and McBride, 2001). The following chapters will address the mode of action, regulation, structure, function, and cellular relevance of this remarkable protein family and of the effectors regulating the activity of small GTPases. These GTPases act as molecular switches and their activities are controlled by a large number of regulatory molecules that affect either GTP loading (guanine nucleotide exchange

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Table 5.2 The Rab GTPase family: Nomenclature, subcellular localization, and possible functions Rab/Swiss-Prot entry

Rab1A (P62820) Rab1B (Q9H0U4) Rab2A (P61019)

Synonym

Localization

Involved in

–

Golgi, ER

ER–golgi trafficking

–

Golgi, ER

ER–golgi trafficking

–

Golgi, ER, ER–golgi intermediate compartment membrane, melanosomes Cell membrane, golgi, ER Cell membrane

ER–golgi trafficking

Rab2B – (Q8WUD1) Rab3A – (P20336)

ER–golgi trafficking

Rab3B (P20337) Rab3C (Q96E17) Rab3D (O95716)

–

Cell membrane

Exocytosis, neurotransmitter release Protein transport

–

Cell membrane

Protein transport

Rab16

Cell membrane

Rab4A (P20338) Rab4B (P61018) Rab5A (P20339)

–

Cell membrane

Protein transport, regulated exocytosis Protein transport

–

Cell membrane

Protein transport

–

Fusion of plasma membrane and EE

Rab5B (P61020)

–

Rab5C (P51148)

Rab5L

Rab6A (P20340) Rab6A0 (P20340)

–

Cell membrane, EE, melanosomes Cell membrane, EE, melanosomes Cell membrane, EE, melanosomes Golgi membrane

Golgi–ER trafficking

Rab6C

Golgi membrane

Golgi–ER trafficking

Protein transport

Protein transport

x

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Table 5.2 (continued) Rab/Swiss-Prot entry

Synonym

Rab6B – (Q9NRW1) Rab7A – (P51149)

Localization

Involved in

Golgi membrane

Golgi–ER trafficking Late endocytic transport, maturation of phagosomes

Vesicular trafficking, neurotransmitter release Vesicular trafficking, neurotransmitter release Protein transport between endosomes and trans-golgi network Protein transport between endosomes and trans-golgi network Vesicular trafficking, neurotransmitter release Endosomal trafficking Transferrin receptor recycling Polarized transport, assembly/activity of tight junctions

Rab7B (Q96AH8) Rab8A (P61006)

–

LE, lysosomes, cytoplasmic vesicles, phagosomes, melanosomes Lysosomes

–

Cell membrane

Rab8B (Q92930)

–

Cell membrane

Rab9A (P51151)

–

Cell membrane

Rab9B (Q9NP90)

Rab9L

Cell membrane

Rab10 (P61026)

–

Cell membrane

Rab11A (P62491) Rab11B (Q15907) Rab13 (P51153)

–

Cell membrane, RE Cell membrane

Rab14 (P61106)

–

Cell junction, tight junction, cell membrane, cytoplasmic vesicles Cell membrane

Rab15 (P59190)

–

Cell membrane

– –

Protein transport

Vesicular trafficking, neurotransmitter release Neurotransmitter release (continued)

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Table 5.2 (continued) Rab/Swiss-Prot entry

Synonym

Localization

Involved in

Rab17 (Q9H0T7) Rab18 (Q9NP72)

–

Cell membrane

–

Cell membrane

Rab19 (A4D1S5) Rab20 (Q9NX57) Rab21 (Q9UL25)

Rab19B

Cell membrane

Transcellular transport Apical endocytosis/ recycling, transport between plasma membrane and EE Unknown

–

Golgi

–

Golgi membrane

Rab22A (Q9UL26)

–

Endosome membrane

Rab23 (Q9ULC3) Rab24 (Q969Q5)

–

Cell membrane

Rab25 (P57735)

–

ER–golgi intermediate compartment, LE Cell membrane

Rab26 (Q9ULW5) Rab27A (P51159) Rab27B (O00194)

–

Secretory granules

– –

Cell membrane, melanosomes Cell membrane

Rab28 (P51157) Rab28L (P51157-2) Rab30 (Q15771)

Rab26

Cell membrane

Targeting uroplakins to urothelial apical membranes Unknown

–

Cell membrane

Unknown

–

Cell membrane

Unknown

Apical endocytosis/ recycling Cell adhesion, endosomal trafficking Sorting of transferrin to recycling endosomes Unknown Autophagy-related processes

Apical recycling and/ or regulating transcytotic pathways Exocrine secretion Exocytosis

x

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Table 5.2 (continued) Rab/Swiss-Prot entry

Synonym

Localization

Involved in

Rab31 (Q13636) Rab32 (Q13637)

Rab22B –

TGN, cell membrane TGN, melanosomes (mitochondria?)

Rab33A (Q14088) Rab33B (Q9H082) Rab34 (Q9BZG1)

–

Golgi membrane

–

Golgi membrane

Rab39

Golgi

Rab35 (Q15286)

Rab1C

Rab36 (O95755)

–

Plasma membrane, endocytic compartment, melanosomes Golgi membrane

Anterograde transport TGN to melanosomes trafficking (Wasmeier et al. (2006)) (mitochondrial fission?) Autophagosome formation Autophagosome formation Redistribution of lysosomes to the peri-Golgi region Regulating endocytic pathways

Rab37 (Q96AX2) Rab38 (P57729)

–

Secretory granules

–

Melanosomes

Rab40A (Q8WXH6) Rab40B (Q12829) Rab40C (Q96S21) Rab41 (Q5JT25) Rab43 (Q86YS6) Rab44 (Q7Z6P3) Rab45 (Q8IZ41)

–

Cell membrane

Melanosomal transport and docking, sorting of TYRP1 Unknown

–

Cell membrane

Unknown

–

Cell membrane

Unknown

–

Unknown

Unknown

Rab41, Rab11B –

ER, golgi

ER–golgi trafficking

Cell membrane

Unknown

RASEF

Perinuclear region

Unknown

Protein transport, vesicular trafficking Unknown

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Partners of small GTPases effectors

GAP (GTPase activating protein)

GEP (GDP/GTP exchange proteins)

GDI (GDP dissociation inhibitor) Ras p21 GAP (100/120 kD) Rho GAP

(p50RhoGAP)

GDS (GDP dissociation stimulator)

Ypt 1 GAP

Ras p21 GIP

Ras p21 GDS (40 kD) Rho GDI a /b (27 kD)

Rab GDS (53 kD)

Rab 11 GDI (54 kD)

Rho GDS

Rab3A GDI (54 kD)

Ran GDS

Rab1 GAP Rab3A GAP

GIP (GTPase inhibiting protein)

Rap1 GDS

Rgd1 (Rho3/4 in yeast) Rac GAP

Figure 5.1 Effectors of small GTPases. A selection of various effectors of small GTPases in mammals and yeast is summarized.

factors or GEFs) or GTP hydrolysis (GTPase activating proteins or GAPs). A choice of effectors/regulators of Ras related GTPases in mammals and yeast cells is summarized in Fig. 5.1. In general, the two different kinds of GTPase regulators include: the proteins that act as activators and those that are able to inactivate the small G-proteins (Seabra and Wasmeier, 2004). The GTPase inhibiting proteins (GIP) keep the protein in its GTP-bound active state, whereas the GDP dissociation stimulator (GDS) works as an activator by supporting the exchange from GDP to GTP. The GAP and the GDP dissociation inhibitor (GDI) serve as inactivating effectors by stimulating the intrinsic GTPase activity, respectively, supporting the maintenance of the GDP-bound state.

2. Rab Proteins 2.1. Rab family Rab GTPases are key players that determine organelle identity and operate at the center of fusion events. They act as switches and are probably linked to each other via their effectors by coordinating cellular transport processes.

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Within this review, we focus on selected examples, the Rab1 and Rab6 proteins, which highlight these issues. The small GTPase family of Rab proteins plays important roles in various aspects of membrane traffic control between intracellular compartments. The Rab proteins constitute the largest subfamily of the Ras superfamily of small GTPases, with more than 60 members encoded by the human genome and until recently 11 members in yeast, the so-called Ypt (yeast protein transport) molecules (Deneka et al., 2003; Jordens et al., 2005; Novick and Zerial, 1997; Pereira-Leal and Seabra, 2001; Pfeffer and Aivazian, 2004; Schwartz et al., 2007). Very recently, a new GTPase in yeast was detected. The entire ORF encoded a Ras-like protein of 205 amino acids and the sequence was deposited with the EMBL databank under the accession number AM696207 and called Ypt12 (Neuhaus et al., 2007). By interacting with a wide variety of effectors, they play a key regulatory function in coordinating the consecutive stages of membrane transport, including the formation of transport carriers and their tethering/docking to target membranes (Chavrier and Goud, 1999; Zerial and McBride, 2001). Several roles displayed by Rab and Rab effector molecules in cellular membrane transport processes are discussed: (1) (2) (3) (4)

Sorting function in vesicle formation. Motor and motor adaptors in intracellular transport processes. Vesicle tethering. Membrane fusion.

The exact cellular localization and tissue-specific expression profile of most of the Rab proteins, so far, is still unknown. In Fig. 5.2, the cellular localization of some prominent members of the endo- and exocytotic Rab family of proteins is summarized (Fig. 5.2; Table 5.2). For instance, Rab33, Rab6, and Rab2, which are located at the Golgi apparatus are involved in exocytotic processes by promoting the Golgi–ER transport (Echard et al., 2000; Valsdottir et al., 2001). Exocytosis of secretory granules and lysosomes is controlled by Rab27 (Blott and Griffiths, 2002; Izumi et al., 2003). Rab1 and Rab2, on the other hand, regulate the anterograde transport from the ER to the Golgi apparatus (Tisdale et al., 1992). Rab4 controls an early sorting event on the endocytic pathway and Rab5, Rab7, and Rab11 also participate in the endosomal pathway (Bucci et al., 2000; Casanova et al., 1999; Mukhopadhyay et al., 1997; Press et al., 1998; Ren et al., 1998; van der Sluijs et al., 1992). For example, Rab5 supports the fusion of endocytotic vesicles and the formation and transport of early endosomes, whereas Rab11 is involved in the endocytotic recycling processes (Bucci et al., 2000; Feng et al., 1995; Gorvel et al., 1991; Nielsen et al., 1999; Novick and Zerial, 1997). Also Rab22 plays a role in the endosomal pathways by controlling the fusion of early endosomes (Kauppi et al., 2002).

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Nucleus Rab24

Autophagosome Melanosome Rab27 Lysosome

Rab1/ Ypt1 Rab2

LE Rab7/ Ypt7

CGN

Rab9

Rab22 Rab4

ER Rab33 Rab6/ Ypt6

EE Rab25

MTOC

Rab5

RE CCV

Rab11/ Ypt11

TGN

Rab5

Figure 5.2 Localization of endo- and exocytotic Rab proteins. Small Rab/Ypt GTPases act in different compartments inside the cell and are able to regulate diverse transport processes. CCV, clathrin coated vesicle; CGN, cis-golgi network; EE, early endosome; ER, endoplasmatic reticulum; LE, late endosome; MTOC, microtubule organizing center; RE, recycling endosome; TGN, trans-golgi network.

Like Rab11, Rab9 regulates receptor recycling processes by controlling the transport of its cargo back to the trans-Golgi network (Lombardi et al., 1993). Rab24 is involved in inducing autophagy (Munafo´ and Colombo, 2002).

2.2. GTPase cycle of Rab proteins Rab/Ypt proteins cycle between an active, GTP-bound, and an inactive, GDP-bound state (Fig. 5.3; Stenmark et al., 1994). Once activated, Rab GTPases are able to interact with downstream effectors. The guanine nucleotide-binding status of Rab proteins is predominantly modulated by three different classes of regulatory proteins, namely the GEFs, GAPs, and GDIs. The Rab GEF proteins regulate Rab GTPase activity by promoting GDP–GTP exchange. Binding of GEF to the Rab protein causes a decrease in Rabs’ affinity for GDP (Horiuchi et al., 1997; Jones et al., 2000). The GAP proteins serve as negative regulators by increasing the intrinsic GTPase activity of Rab proteins and therefore inducing the conversion of the active GTP-bound state to the inactive GDP-bound state (Clabecq et al., 2000; Haas et al., 2005; Strom et al., 1993; Will and Gallwitz, 2001). The GDI proteins also act as negative regulators by blocking GDP dissociation via interactions with the isoprenylated carboxyterminus of Rab proteins (DerMardirossian and Bokoch, 2005; Pfeffer and Aivazian, 2004). Newly synthesized, the Rab proteins are GDP-bound and associated with a Rab escort protein (REP). They have to be geranylgeranylated at either one

P

GAP

GDI

GDP GTP REP

GGTase

GEF

GGTase Targeting

GDF Active Rab

Inactive Rab

REP

GDI

GGTase

GDF

Figure 5.3 The GTPase cycle. Rab/Ypt GTPases cycle between a GTP bound active (blue square) and a GDP bound inactive (blue circle) state. After their synthesis Rab proteins are geranylgeranylated by the geranylgeranyltransferase (GGTase) and delivered to their target membrane by the Rab escort protein (REP). To activate Rab, the guanine-nucleotide exchange factor (GEF) replaces GDP with GTP. The inactivation is catalyzed by a GTPase activating protein (GAP). The GDI not only keeps the Rab protein in this state by stabilizing it, but also transports it to the cytoplasm and back to the membrane. Because this interaction is relatively strong, the release of the GTPase from the GDI has to be supported by another protein: the GDI-displacement factor (GDF).

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or two cysteine residues located at or near its carboxy terminus (see Section 2.3). This prenylation is necessary for anchoring the Rab protein in membranes and is carried out by the geranylgeranyltransferase (GGTase). After prenylation the Rab/Ypt protein is delivered to its target membranes by the REP (Casey and Seabra, 1996). To activate the Rab protein, GDP has to be exchanged against GTP. This replacement is catalyzed by GEF (Ali and Seabra, 2005; Alory and Balch, 2003). Burton and colleagues discovered MSS4 as the first GEF, which affects Sec4 (Burton et al., 1993). After activation, the Rab can fulfil its function by interacting with effector proteins. The Rab protein can be switched off by GTP hydrolysis, which is an intrinsically very slow process and thus needs to be accelerated by a GAP (Vetter and Wittinghofer, 2001). The first Ypt GAP that was discovered is Gyp6, which activates the intrinsic GTPase activity of Ypt6 (Strom et al., 1993). The GDI inhibits the exchange from GDP to GTP and thus keeps the Rab protein in its inactive state. It is also responsible for transporting Rab to the cytoplasm and back to the membrane. GDI1p was the first protein to be found which acts as GDI for Sec4p (Garrett et al., 1994). GDIs and REPs are structurally closely related and act in a similar manner. They recognize the GDP-bound conformation of the Rab, bind the C-terminal region (including the lipid-modifications) in an extended conformation via hydrophobic residues and thus assure the solubility of the GTPase (Rak et al., 2003, 2004). The exact mechanisms of REP/GDI–Rab interactions have been summarized in an excellent review by Seabra and Wasmeier (Seabra and Wasmeier, 2004). After the GDP-bound Rab has been escorted to the membrane again, the GDI is released with the help of a GDI displacement factor (GDF) and activation via GEF completes the cycle (Ali and Seabra, 2005). For studying the functions of Rab/Ypt proteins, a set of dominant negative and constitutively active mutants was created to simulate the active or inactive state of the GTPase. Table 5.3 shows an assortment of Rab/Ypt mutants, which were used for the experiments described below.

2.3. Structure and phylogeny of Rab GTPases The general structure of Rab GTPases follows the structure of all small GTPases of the Ras superfamiliy described above. The core fold consists of six b-sheets and five a-helices. Within these structures, the nucleotide- and phosphate/Mg2+-binding regions are located. Like in all small GTPases, the major conformational changes upon GTP binding and hydrolysis take place within the switch I and switch II regions. Thus, the switch regions represent the structural elements displaying the on/off-state of the protein (Vetter and Wittinghofer, 2001). In Fig. 5.4A, the approximate positions of these structural features are highlighted within a sequence alignment of multiple Rab proteins.

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Table 5.3 Active and inactive mutants of Rab1/6/11 and their yeast homologues Ypt1/Ypt6/Ypt11

Rab/accession numbers

Constitutively active mutants

Dominant negative mutants

Rab1A (human) (NM_004161) Rab1B (rat) (X13905) Rab6A (human) (NM_198896) Rab6B (human) (NM_016577) Rab11A (human) (Image clone: 151446) Ypt1 (yeast) (X00209) Ypt6 (yeast) (X59598) Ypt11/Ypt32 (yeast) (U33754)

Q70L Q67L/R Q72L/R Q72L/R S20V Q67L Q69L S22V

S25N S22N T27N T27N K24R S22N T24N K26R

Further, all Rab proteins contain an unfolded hypervariable carboxyterminal domain. These carboxyterminal domains are highly divergent between all Rab sequences and are supposed to have an impact on targeting of a Rab to its specific compartment (Chavrier et al., 1991; Pfeffer, 2005). From the alignment and the cladogramm shown in Fig. 5.4A and B it becomes obvious that even closely related Rab proteins differ significantly within their carboxyterminal sequences. The hypervariable domains end with the prenylation site, which contains two cystein residues in one of the following combinations: XXXCC, XXCXX, XCCXX, CCXXX, or XXCXC (Pereira-Leal and Seabra, 2000). Some attempts were made to combine structural and sequence data with functional features and to solve the mechanisms underlying the interaction of Rab proteins and their effectors. Ostermeier and Brunger (1999) published their structural analyses of the interaction of Rab3A with its effector Rabphilin-3A. Based on these data they defined the Rab complementarity-determining region (RabCDR), a pocket that, apart from the switch regions, interacts with effector molecules and thus could be responsible for the specificity of effector binding. One year later, Pereira-Leal and Seabra, in a primary sequence analysis, defined five sequence regions that discriminate Rabs from other small GTPases, the so-called Rab family or RabF regions. Additionally, they spotted four regions that are specific for certain subfamilies of Rab proteins, RabSF1–RabSF4. Interestingly, the RabSF1, RabSF3, and RabSF4 regions mainly overlap with the sequence regions that form the RabCDR. RabSF2 is adjacent to the switch I region and thus could be responsible for further specific effector interactions. In a subsequent study, the RabF motifs were shown to be responsible for the interaction with general regulators like the REP (Pereira-Leal et al., 2003). In the alignment shown in Fig. 5.4A, the RabF and RabSF regions are denoted according to the consensus sequences described by Pereira-Leal and Seabra (2000).

248 A

a1

b1

b5

a4

b2

b6

b3

a5

Figure 5.4 Continued

a2

b4

a3

B

Ha-Ras

Ypt1 Rab1A Rab1B Ypt11 Rab11A Rab11B Ypt6 Rab6B Rab6A Rab6A‘

Figure 5.4 Continued

249

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Angelika Barnekow et al.

Figure 5.4 Structure and phylogeny of Rab GTPases. (A) Alignment of human Rab1, 6 and 11, and their yeast homologues; Ha-Ras is aligned as reference; the alignment was produced using ClustalW and manually adjusted within the C-terminal regions so that the prenylation sites within each group of homologous proteins match with each other. Regions that form a-helices and b-sheets, the switch regions as well as nucleotide- and phosphate/Mg2+-binding sites (G1–G3 and PM1–PM3, marked with asterisks) are denoted above the sequences. RabF and RabSF regions are marked within the alignment in red and orange, respectively, and are labelled below the sequence. (B) Schematic representation of the phylogenetic relationship between the Rab GTPases aligned in Fig. 5.4A. The tree was calculated using ClustalW and displayed using treeview with

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Merithew et al. identified another important component for a Rab/effector interaction, an invariant hydrophobic triad consisting of a phenylalanine, a tryptophan, and a tyrosine residue, for example, F58-W75-Y90 in Rab5, F59W76-Y91 in Rab3A, or F50-W67-Y82 in Rab6A, respectively (Merithew et al., 2001; Schweizer Burguete et al., 2008). These amino acids are located within the switch regions and have been shown to be directly involved in contact with the effector molecule (Schweizer Burguete et al., 2008). Although this triad is highly conserved, the conformation of their bulky side-chains can differ significantly depending on the adjacent sequences. Thus, the variable surrounding of the switches can induce a structural plasticity within these highly conserved regions. This gives rise to additional specificity in the interactions between Rab subfamilies and their effectors. Taken together, these data provide a nice model for Rab/effector interactions: The interaction of a Rab protein with its effectors generally depends on the active or inactive state of the Rab protein, represented by the conformation of its switch regions; RabF regions define conserved binding surfaces for general effectors like REP or GDI; specificity of effector binding to one Rab or a defined subgroup is achieved by variable subfamily specific regions that are the basis for additional interacting elements like the RabCDR and the conformation of the invariant hydrophobic triad within the switches. Pereira-Leal and Seabra published an evolutionary analysis of complete Rab GTPase sets from six different organisms (Pereira-Leal and Seabra, 2001). This study revealed that Rab proteins can be grouped into eight interspecies-specific functional groups that show similarity not only on a sequence level but also in terms of cellular localization and/or function. Most, but not all Rab proteins can be assigned to one of these subgroups. These groups are most likely to reflect a common ancestral origin, giving rise to the hypothesis that an ancestral ‘‘basic’’ set of Rab proteins developed at an early point within evolution, that is, before the separation of plants, fungi, and animals. The development of more proteins and isoforms happened later within the species and subgroups. In 1997, the complete genome sequencing of Saccharomyces cerevisiae revealed a total number of 11 Ypt proteins (Lazar et al., 1997). Recently, a new Ypt gene was detected, the Ypt12 and the sequence was deposited with the EMBL databank under Ha-ras defined as outgroup. The lengths of the branches do not correspond to absolute phylogenetic distances, but rather show the degree of relationship on the basis of primary sequence changes. (C) Localization of endogenous Rab6A and transiently transfected EGFP-Ypt6 Q69L in HeLa cells. Cells were transfected using Polyfect and incubated for 36 h. Staining of endogenous Rab6A was performed using the mouse antibody Mab 5B10 (1:50) and a Cy3-coupled anti-mouse secondary antibody (1:1000). Nuclei were stained with DAPI. Fluorescence pictures were acquired and processed using an Olympus BX-61 microscope with a 100 objective and the Cell^P software. Rab6A Cy3 signals are shown in the upper panel, EGFP-Ypt6 Q69L in the middle, and an overlay in the lower panel.

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the accession number AM696207. Multicellular organisms today generally contain a higher number of Rab isoforms that are often tissue- or cell-type specifically expressed. Many of these isoforms occurred as a result of gene or exon duplications, for example, Rab6A and Rab6A´ (Del Nery et al., 2006; Echard et al., 2000). It has been proposed that this enlargement of the Rab GTPase inventory was necessary to fulfil the requirements of the increasingly complex cell physiology and membrane trafficking events in multicellular organisms (Bock et al., 2001). This development of the Rab/Ypt family can already be retraced from the relatively small amount of Rab/Ypt sequences, the phylogeny of which is depicted in Fig. 5.4B. The cladogram shows the phylogenetic relationship between all human isoforms of Rab 1, 6, and 11 and their yeast counterparts. The proteins cluster together on the branches of the tree according to their sequence similarity, the nodes represent a common ancestral origin. It becomes obvious that the proteins do not cluster according to their species affiliation into human and yeast groups of proteins, but in fact, homologous proteins that belong to the same functional group cluster together despite their origin. Human Rab6 isoforms and yeast Ypt6 form a branch that can clearly be separated from the other groups, whereas the Rab1/Ypt1 and Rab11/Ypt11 groups form a common branch and are thus more closely related to each other than to the Rab6 group. Further, the tree displays that the split up of each Rab protein and its corresponding Ypt took place before the occurrence of the multiple Rab isoforms. The common origin of Rab and Ypt proteins becomes obvious not only on a sequence basis, but also in terms of functional similarities. In mammalian cells, both Rab1A and B are involved in anterograde ER to Golgi as well as intra-Golgi-transport events (Nuoffer et al., 1994; Plutner et al., 1991; Saraste et al., 1995; Tisdale et al., 1992). Rab6 isoforms play a role in retrograde Golgi to ER and intra-Golgi-transport and recently, Rab6 has also been proposed to be involved in the transport of exocytotic carriers (Grigoriev et al., 2007; Martinez et al., 1994, 1997; Matanis et al., 2002; for a detailed discussion of Rab1 and Rab6 functions see Chapter 3). In yeast, Ypt1 can be functionally replaced by its mouse homologue and the tethering of ER-derived vesicles to Golgi membranes has been shown to be Ypt1-dependent (Cao et al., 1998; Haubruck et al., 1989). For Ypt6, an important role in retrograde Golgi transport processes has been described (Luo and Gallwitz, 2003). Further, immunofluorescence microscopy experiments performed by our group indicate a similar localization pattern for Rab6A and Ypt6 Q69L in human HeLa cells (Fig. 5.4C). Although there are differences in the precise site of action of Ypt11 in yeast compared to mammalian Rab11, it was shown that yeast Ypt11, expressed in mammalian cells, is targeted to Rab11A-positive compartments (Kail et al., 2005). Furthermore, transport assays displayed that Ypt11 is functionally active in the membrane recycling systems of mammalian cells. Another interesting aspect regarding the evolution of Rab/Ypt proteins

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is the interaction with their general effectors. As these interactions can be very specific and highly conserved, coevolution of Rab proteins and their effectors must have taken place.

2.4. Rab GTPases and malignancies Several studies on vesicular trafficking in secretory and endocytic pathways have linked dysfunction of Rab GTPases to human diseases. Mutations in general regulators of Rab activity were shown to cause malignancies. Deficiencies in the Rab prenylation machinery can lead to different diseases like Choroideremia, Hermansky–Pudlak syndrome (HPS), or Griscelli syndrome (GS) (Pereira-Leal et al., 2001). Choroideremia is an X-chromosomelinked disease that leads to the degeneration of the choriocapillaris, the retinal pigment epithelium and the photoreceptor layer in the eye. Choroideremia patients have loss of function mutations in the Rab escort protein, REP1, which is identical to CHM, the gene product, which is defective in choroideremia. CHM/REP proteins are evolutionarily related to the GDI family of proteins (Alory and Balch, 2001). The HPS, an autosomal recessive disorder, which is characterized by oculocutaneous albinisms and hemorrhagic episodes due to platelet storage pool deficiency, possesses a splice-site mutation in the a-subunit of geranylgeranyltransferase (RGGT), which results in hypopigmentation and platelet dysfunction (Hermansky and Pudlak, 1959; Pereira-Leal et al., 2001). The small GTPase Rab38 is also implicated in the pathogenesis of this disease (Di Petro and Dell’Angelica, 2005; Osanai and Voelker, 2008). Rab38 was identified as a protein that is predominantly expressed in pigmented melanocytes and retinal pigment epithelium cells. In addition, Rab38 mRNA is expressed in 80–90% of melanoma, but rarely in nonmelanocytic malignancies ( Ja¨ger et al., 2000). Mutations in the Rab GDI-a, which is postulated to extract GDP-bound Rab proteins from target membranes and to maintain them in the GDPbound state, are involved in X-linked mental retardation (Alory and Balch, 2003; D’Adamo et al., 1998). Rab GDI-a is predominantly expressed in brain and seems to regulate locally abundant Rab proteins like the Rab3 isoforms (Alory and Balch, 2003). Evidence is accumulating that Rab3 plays a key role in neurotransmitter release and synaptic plasticity. Mutations in the Rab3 GAP were found to cause Warburg Micro syndrome and Martsolf syndrome (Sakane et al., 2008). Both malignancies lead to mental retardation. In addition, mutations in the Rab27A gene also result in hypopigmentation due to clustering of melanosomes in GS, caused by defects in melanosome transport and loss of cytotoxic killing activity in T cells (Pereira-Leal et al., 2001; Seabra et al., 2002). GS is a rare autosomal recessive disorder characterized by pigment dilution of hair and an uncontrolled T-cell and macrophage activation syndrome known as hemophagocytic syndrome (Klein et al., 1994).

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GS patients exhibit loss of cytotoxic T lymphocytes (CTL) killing activity. The majority of GS patients, which exhibit defects mainly in two types of lysosome-related organelles, melanosomes in melanocytes and lytic granules in CTLs, have a defect in the Rab27A gene (Barral et al., 2002). The X-linked disorder oculocerebrorenal syndrome of Lowe is caused by mutation in the OCRL1 protein. OCRL1 encodes an inositol polyphosphate 5-phosphatase and is localized to the Golgi apparatus and early endosomes (Hyvola et al., 2006). Several small GTPases interact with the OCRL1 protein. The strongest signals were obtained with Rab1, Rab5, and Rab6 and it was concluded that Rab proteins play a dual role in the regulation of OCRL1 by firstly targeting this protein to the Golgi apparatus and endosomes, and secondly, by directly stimulating the 50 -phosphatase activity of OCRL1 after membrane recruitment (Hyvola et al., 2006). Another involvement of the Rab1 and Rab6 GTPases in human malignancies has been reported by Wu et al. They found that Rab1, Rab4, and Rab6, but not Rab3 and Rab5, are upregulated in a dilated cardiomyopathy model overexpressing ß2-adrenergic receptors (Wu et al., 2001). Rab1A overexpression caused cardiac hypertrophy and the authors concluded that increased expression of Rab1 in myocardium distorts subcellular localization of proteins and is sufficient to induce cardiac hypertrophy and failure (Wu et al., 2001). Many Rab GTPases regulate the intracellular transport and the cellular localization of the cystic fibrosis transmembrane conductance regulator (CFTR), a Cl–-selective ion channel, which belongs to the ATP-binding cassette (ABC) transporter superfamily (Guggino and Stanton, 2006). Mutations in ABC genes have been linked to many genetic diseases, including cystic fibrosis, which is characterized by an exocrine pancreatic insufficiency, an increase in sweat NaCl concentration, male infertility, and airway disease. It was shown that Rab5 regulates the transport of CFTR from the plasma membrane to early endosomes. The recycling back to the plasma membrane is controlled by Rab11. Rab7 controls the trafficking of CFTR from early endosomes to late endosomes. Rab9 can transport CFTR from late endosomes to the trans-Golgi. From their findings, the authors conclude that small GTPases of the Rab family are potential targets that might be used to develop new drugs for the treatment of cystic fibrosis and secretory diarrhoea. Also for neuronal malignancies, an involvement of Rab GTPases is discussed. Devon et al. reported that Als-2-deficient mice show an altered Rab5dependent endosomal transport (Devon et al., 2006). ALS2 is an autosomal recessive form of a motor neuron disease caused by loss of function of alsin, the protein encoded by ALS2. This protein has been characterized as GEF for Rab5, which is known to be an important factor in endocytosis and early trafficking of signaling molecules. In neurons, the Rab5 GTPase mediates endocytotic transport, which is essential for presynaptic transmission and for

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the internalization of a-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors. Its interaction with alsin has been suggested to modulate the signal transduction of neurotrophic factors like insulin-like growth factor (IGF1) and brain-derived neurotrophic factor (BDNF) receptors. Recently, Huntingtin (Htt) and Huntingtin associated protein (HAP40) were described as a novel Rab5 effector complex that regulates endosome motility (Pal et al., 2006). In Huntington disease (HD), HAP40 protein expression was increased leading to an elevated recruitement of Htt onto Rab5-positive early endosomes that drastically reduce their mobility suggesting a mechanism by which impaired Rab5-mediated transport of neurotrophic factors may be a key event of the pathological process leading to neurodegeneration in HD (Pal et al., 2008). The Rab5 protein, especially the Rab5B protein, may provide a link between neurodegenerative disease, neuroprotection, and synaptic plasticity and therefore possibly being a useful target for drug development (Baskys et al., 2007). Another neurodegenerative disease, the Parkinson’s disease (PD), is an age-associated movement disorder. Gene mutations within the a-synuclein gene are major genetic defects known in the familial juvenile onset of PD. The a-synuclein protein blocks endoplasmic reticulum-to-Golgi-transport. It has been shown, that overexpression of the Rab1 GTPase can suppress the a-synuclein toxicity and can rescue neuron loss in a-synuclein-induced neurodegeneration (Chua and Tang, 2006). Another Rab GTPase, the Rab6 protein, has also been shown to be involved in a neurodegenerative malignancy, the Alzheimer’s disease (McConlogue et al., 1996; Scheper et al., 2007; Teber et al., 2005; see also paragraph 3). There are several investigations, which point to an emerging role of Rab GTPases in cancer (Cheng et al., 2005). One example for the involvement of Rab GTPases in tumor formation is the human Rab2B, which was shown to be overexpressed in colon carcinoma CX1 (Ni et al., 2002). The observation that the Rab2B gene is extremely high expressed in colon adenocarcinoma CX-1 implies a close relationship with colon tumors. Recently, it was shown, that Rab25 is amplified and overexpressed in ovarian and breast cancer (Cheng et al., 2004, 2005). The increase in Rab25 expression was stage-dependent suggesting a role in tumor progression and aggressiveness. The authors discuss that Rab25 is not sufficient to induce tumor formation but rather increases the rates of tumor growth and aggressiveness in already transformed cells (Cheng et al., 2004). For the Rab25 GTPase an upregulation in relation to tumorigenicity has also been reported in prostate cancer (Calvo et al., 2002). Also Rab27A in context with its effector JFC1/Slp1 are important regulators of exocytosis processes in prostate carcinoma cells (Catz, 2008). An aberrant expression of the Rab1A gene has been described for human tongue cancer (Shimada et al., 2005). The authors suggest that the Rab1A protein could be a potential biomarker of tongue carcinogenesis. Several other small GTPases of the Rab subfamily are discussed to be involved in hepatocellular carcinomas. Rab1B, Rab4B, Rab10, Rab22A, Rab24, and

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Rab25 were shown to be upregulated in most of the 11 hepatocellular carcinomas tested (He et al., 2002; Yao et al., 2003). For Rab31, Kotsch et al. could show a correlation between the malignancy of breast tumors and the degree of Rab31 mRNA expression (Kotsch et al., 2008). High levels of Rab31 were associated with worse outcome of patients for distant metastasis-free survival. Despite recent progress in investigating the role of Rab proteins in human cancer development and/or progression, our knowledge is still very limited and further studies are needed to evaluate a concrete function of Rab GTPases in tumorigenesis. Several reports address the potential involvement of Rab GTPases in infectious diseases caused by protozoa, bacteria, and viruses. In Listeria monocytogenes infected macrophages, the small GTPase Rab5A is overexpressed and accelerates the intracellular degradation of the bacteria, while suppression of Rab5A results in enhanced Listera survival (Alvarez-Dominguez and Stahl, 1999). Another example for altered expression profiles of Rab GTPases in bacterial caused diseases is the downregulation of Rab33A in patients with Tuberculosis (TB), which remains a major human health threat caused by Mycobacterium tuberculosis ( Jacobsen et al., 2005). Differential gene expression analyses using microarrays from patients with TB revealed three differentially expressed Rab genes. Rab33A was down-regulated, whereas Rab13 and Rab24 were upregulated. The studies identified Rab33A as a T cell regulatory molecule in TB being involved in the process of the TB disease. Chlamydiae are obligate intracelluar bacteria that cause bacterially acquired sexually transmitted disease and preventable blindness worldwide. Rzomp et al. succeeded in showing that Inc CT229, one of the chlamydial inclusion membrane proteins (Incs) interacts with Rab4A and recruits the small GTPase to the inclusion membrane (Rzomp et al., 2006). The interaction was detected using the yeast two-hybrid system and confirmed by GST pulldown experiments and colocalization studies (Rzomp et al., 2006). Another link between small GTPases of the Rab subfamily and pathogens was reported by Batista et al. (2006). The authors found that the protozoan Trypanosoma cruzi, which causes the Chagas disease, alters the expression of Rab GTPases. T. cruzi-infected cardiomyocytes displayed a downregulation of Rab7 and Rab11, whereas the expression of Rab5A was maintained suggesting that T. cruzi impairs the endocytosis in mouse cardiocytes (Batista et al., 2006). Rab proteins have already been shown to be implicated in the life cycles of various viruses (Brass et al., 2008; Krishnan et al., 2007; Sklan et al., 2007a; Stone et al., 2007; Vonderheit and Helenius, 2005). Recently, an involvement of the Rab1 protein in Hepatitis C Virus (HCV) replication has been published (Sklan et al., 2007b). It was shown, that TBC1D20, the first known GAP for Rab1, mediates HCV replication. Furthermore, Rab1 depletion significantly decreased the amount of HCV RNA suggesting that this GTPase may also be involved in HCV replication (Sklan et al., 2007b).

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3. Small GTPases Rab1, Rab6, and Their Effectors The following paragraph summarizes data on Rab1 and Rab6 as prominent examples for small GTPases in more detail. With the help of constitutively active and inactive mutants, we and others were able to study activity-dependent interactions of the Rab1 and Rab6 GTPases. The experimental approach often starts with an extensive yeast two hybrid (YTH) screen using Rab-specific cDNA coding for the active or inactive Rab Protein as bait construct, which is followed by either GST pull down analyses or coimmunoprecipitations to verify the specificity of potential Rab interacting partners isolated via the YTH screen (Kail and Barnekow, 2008; Monier and Goud, 2005). Colocalization studies applying immunofluorescence microscopy or cell fractionation experiments using differential or density gradient centrifugation complete the investigations. As examples for mammalian RabGTPase effectors we describe the attempts, which have been made to detect and isolate specific partners of the Rab1 and Rab6 GTPase. The two isoforms of Rab1, Rab1A and Rab1B, are located to the endoplasmic reticulum (ER), pre-Golgi intermediates, and the Golgi stack. Both proteins have been suggested to regulate anterograde transport of cargo between the ER and the Golgi apparatus (Nuoffer et al., 1994; Sannerud et al., 2006; Saraste and Goud, 2007; Saraste et al., 1995; Tisdale et al., 1992). Rab1 also associates with intermediate compartment (IC) tubules that accumulate in transport-arrested cells and are devoid of cargo proteins raising the possibility that Rab1 proteins function in a retrograde transport process at the ER–Golgi boundary (Palokangas et al., 1998; Sannerud et al., 2003, 2006). To date p115, GM130, golgin-84, the MICAL proteins, Iporin, and Giantin have been identified as partners interacting with the active conformation of Rab1 (Allan et al., 2000; Bayer et al., 2005; Beard et al., 2005; Diao et al., 2003; Fischer et al., 2005; Moyer et al., 2001; Weide et al., 2001, 2003). p115, GM130, Giantin, and golgin-84 belong to a family of coiled-coil proteins, the golgins, playing a role in tethering ER-derived and intra-Golgi vesicles to the Golgi apparatus (Short et al., 2005). Iporin and the MICAL proteins seem to link Golgi associated Rab1 proteins to the intermediate filament cytoskeleton. After constructing and testing various deletion mutants of GM130, Iporin, and MICAL-1A, we were able to map the Rab1 binding domains within the interacting target molecules (Fig. 5.5). Surprisingly, we recently found that the Rab1 binding partner Giantin, a golgin family protein that is attached to Golgi membranes by a putative carboxyterminal membrane anchor, also was able to interact with a second Rab GTPase, the Rab6 molecule (Beard et al., 2005; Rosing et al., 2007; Fig. 5.6). Another example for an interaction of two Rab GTPases, the

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991

0

1239

1516

Iporin

0

370

887 990

GM130 0

836

981

MICAL-1A

Figure 5.5 Rab1 effectors. By constructing and testing various deletion mutants of Rab1 interacting partners in YTH and GST pulldown analyses, we characterized the binding domains of Rab1 to the effector/regulator proteins. The dark grey boxes represent the binding regions.

Figure 5.6 Isoforms of Rab6. A sequence alignment of Rab6A, A0 , and Rab6B is shown. Differences are shown in grey or black, respectively.

Rab6 and the Rab11 proteins, with the same interacting partner is the Rab6-binding protein Rab6IP1 (Miserey-Lenkei et al., 2007). In addition to Rab6, R6IP1, that’s function is required during metaphase and cytokinesis of mitosis, binds to Rab11A in its GTP-bound conformation. It is concluded that Rab11 and Rab6 besides their role in membrane trafficking also function in mitosis and cytokinesis; for example, Rab6A through its interaction with MKpl2 (discussed later) and Rab6A0 through its interaction with GAPCenA and p150Glued participate in pathways involved in the metaphase/anaphase transition (Miserey-Lenkei et al., 2006). Two further cytoplasmic proteins, which were shown to specifically interact with the

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active GTP-bound forms of Rab6A and of Rab6A0 , are the Rab6 interacting proteins 2A and 2B (Rab6IP2A/B/ELKS), which are reported to be ubiquitously expressed (Monier et al., 2002). Rab6 proteins are ubiquitous mammalian proteins primarily located on medial- and trans-Golgi cisternae and on the trans-Golgi network (TGN). In various cell types, Rab6 is also found on post-Golgi vesicles and highly dynamic tubular structures, which move along microtubules (Echard et al., 1998; Martinez et al., 1997; Matanis et al., 2002; Wanschers et al., 2007; White et al., 1999). Rab6 has also been reported as regulator for transport and targeting of exocytotic carriers (Grigoriev et al., 2007). Furthermore, the Rab6 molecule controls the intra-Golgi transport as positive regulator of retrograde transport within the Golgi apparatus. Three isoforms of Rab6 are known in mammalian cells, namely Rab6A, Rab6A0 , and Rab6B (Echard et al., 2000; Goud et al., 1990; Opdam et al., 2000; Fig. 5.6). Rab6A and Rab6A0 represent alternatively spliced isoforms of the same gene differing only in three amino acid residues (Opdam et al., 2000; Fig. 5.6). Whereas Rab6A regulates a coatamer protein I (COPI)-independent pathway between the Golgi apparatus and ER, Rab6A0 has been implicated in Golgi-to-endosomal recycling (Del Nery et al., 2006; Mallard et al., 2002; Young et al., 2005). Rab6B is preferentially expressed in brain and shows a 91% homology to Rab6A (Opdam et al., 2000). So far, a number of interacting partners of Rab6 have been described like GAPCenA, a GTPase-activating protein of Rab6, Rabkinesin6/Rab6KiFL/MKlp2, Rab6-interacting proteins 1/2 (Rab6IP1/2), Bicaudal 1/2, TMF (ARA160), p150Glued, Mint3, Giantin, DNLC2A, PMM1, or GCC185 (Cuif et al., 1999; Echard et al., 1998; Fridmann-Sirkis et al., 2004; Hill et al., 2000; Kail and Barnekow, 2008; Kaufmann et al., 2005; Monier et al., 2002; Rosing et al., 2007; Schweizer Burguete et al., 2008; Teber et al., 2005; Yamane et al., 2007; Figs. 5.7–5.10A and B). There are several links between Rab6 and malignancies. For example, Rab6 has been discussed in context with Alzheimer’s disease by several groups (McConlogue et al., 1996; Scheper et al., 2007; Teber et al., 2005). Very recently, Rab6 has also been reported to be linked to human immunodefiency virus (HIV) entry and productive infection via the regulation of Golgi retrograde pathways (Brass et al., 2008). Rabkinesin-6/RAB6-KIFL, meanwhile referred to as mitotic kinesinlike protein2 (MKlp2), is a vertebrate-specific kinesin-9 family motor protein originally identified as interacting partner of the Rab6 GTPase, which displays a cell cycle-regulated expression pattern being essentially absent from interphase cells and abundant in mitotic cells (Echard et al., 1998; Fontijn et al., 2001; Hill et al., 2000; Neef et al., 2003). MKlp2 belongs to the family of kinesin proteins, which are characterized by a conserved motor domain that binds to microtubules and couples ATP hydrolysis to generate mechanical force. The kinesin family of proteins plays a variety of roles in

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0

262 357

3226

Giantin 524

0

736

1278

Hp150Glued 673

0

803 975

BICD1 630

0

820

BICD2 438

0

624

838

Mint1 749

0 Mint2 0

213

369

576

Mint3 0

262

PMM1 0

96

DLC2A

Figure 5.7 Rab6 effectors. By constructing and testing various deletion mutants of Rab6 interacting partners in YTH and GST pulldown analyses, we characterized the binding domains of Rab6 to the effector/regulator proteins. The dark grey boxes represent the binding regions.

cellular functions, being involved in diverse processes including formation of the mitotic spindle and chromosome partitioning, as well as intracellular movement of organelles and vesicles. The interaction of Rab6 and MKlp2 suggests a role for Rab6 in microtubule-mediated membrane traffic between endosomes and the TGN. A second link between microtubules and the Rab6 proteins comes from the observation that Rab6A, Rab6A0 , and Rab6B can interact with a coiled-coil protein of the trans-Golgi, the Bicaudal 1 and 2 (BicD1 and BicD2) proteins (Kail and Barnekow, 2008; Matanis et al., 2002; Short et al., 2002; Wanschers et al., 2007). This interaction is conserved during evolution and has been recently reported also for the Drosophila melanogaster ortholog of the Rab6 GTPase, Drab6, and the BicD protein (Coutelis and Ephrussi, 2007; Januschke et al., 2007). Human BicD 1 and 2 proteins interact with Rab6 as well as with the dynein–dynactin motor complex (Hoogenraad et al., 2001). In addition, the p150Glued subunit of dynactin can also interact with Rab6A, RabA0 , and Rab6B (Kail and Barnekow, 2008; Short et al., 2002; Table 5.4). Additionally, the authors of another study suggest that dynein could be the motor protein involved in microtubule-dependent Rab6 mediated Golgi-to-ER transport (Young et al., 2005). Recently, we identified dynein light chain 2A (DNLC2A), a human light chain of cytoplasmic dyneins, to be a Rab6 interaction partner using the YTH system ( Jiang et al., 2001; Kail and Barnekow, 2008; Kaufmann et al., 2005; Schroer, 2004). Wanschers et al. reported in 2008 that Rab6 family proteins interact with the dynein light chain protein DYNLRB1 (Wanschers et al., 2008). Dynein consists of two

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20mm

B

20mm

C

20mm

Figure 5.8 Colocalization studies on Rab6A and Mint3 by immunofluorescence. Stable CFP-Rab6A Q72L Polyfect transfected HeLa T-Rex cells were first fixed with 4% paraformaldehyde (PFA)/250 mM Hepes for 15 min on ice, then with 8% PFA/ 250 mM Hepes at room temperature for 30 min and stained with anti Mint3 antibody (1:50, BD Biosiences, Heidelberg, Germany) and with Cy3 coupled anti mouse IgG (1:500, Jackson Immuno Research, Dianova, Hamburg, Germany). For documentation, an Olympus BX61 microscope and the Cell^P software (no neighbour algorithm) were used. Bar: 20 mm.

heavy chains, two intermediate chains, four light intermediate chains, and several light chains, called DYNLRB proteins. In mammalian cells, two light chain isoforms have been identified, DYNLRB1 and DYNLRB2. Applying yeast two hybrid, coimmunoprecipitation, and pull down studies, Wanschers et al. showed that DYNLRB1 specifically interacts with all three Rab6 isoforms and colocalizes at the Golgi (Wanschers et al., 2008).

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Figure 5.9 Localization of endogenous Rab6A and Giantin at the Golgi apparatus. Endogenous Rab6A and Giantin, a coiled-coil golgin, were stained using the Mab 5B10 (1:100) and anti-Giantin (BAbco, USA, 1:1000) primary antibodies. As secondary antibodies, anti-mouse antibodies coupled with Cy3 (1:1000) for Rab6A (panel A) and Alexa488 (1:1000) for Giantin (panel B) were used. In panel C, an overlay of both stainings is shown. It becomes obvious, that Rab6A and giantin colocalize significantly within the Golgi region. Fluorescence pictures were obtained and processed using an Olympus BX-61 microscope with a 100 objective and the Cell^P software.

Another Rab6 interacting protein also containing a short conserved coiled-coil motif, binding the Rab6 isoforms Rab6A, Rab6A0 , and Rab6B, is TMF/Ara160 (TATA element modulatory factor/androgen receptor-coactivator of 160 kDa), which localizes to the Golgi complex and is required for proper Golgi morphology (Fridmann-Sirkis et al., 2004). Recently, it was shown that TMF associates with the budding structures at the tips of the Golgi cisternae and that it modulates two Rab6-dependent retrograde transport processes, a direct endosome-to-TGN and a COPI-in dependent Golgi-to ER route (Yamane et al., 2007).

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A Bait

Prey pACT PMM1 wild type selection

β-gal test

pAS Rab6A wt

+++

++

pAS Rab6A⬘ wt

+++

++

pAS Rab6B wt

++

+

pAS

−

−

B

1

2

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Ha tagged Rab6A

Figure 5.10 Characterization of the interaction between phosphomannomutase 1 (PMM1) and Rab6. (A) Yeast two-hybrid system: Y190 yeast cells were simultaneously transformed with the corresponding bait/prey constructs. The interaction was analyzed by plating the transformed yeast cells on selective media and by ß-galactosidase filter assays (Weide et al., 2001). Selection: his3 selection medium lacking leucine, tryptophan, histidine and supplemented with 30mM 3AT; ß-gal test: ß-galactosidase reporter gene activity; – indicates no growth on selection media or no ß-galactosidase activity; + indicates low growth or low b-galactosidase activity; ++ indicates strong growth on selection media or ß-galactosidase activity; +++ indicates very strong growth on selection media or high b-galactosidase activity. (B). GST pulldown analysis: Escherichia coli BL21 cells were transformed with pGEX encoding GST (glutathione S-transferase) or pGEX42 encoding the fusion protein GST–PMM1 (full length). Expression was induced with 1mM IPTG. Cells were pelleted, resuspended in PBS, 5% Trasylol and sonicated. TritonX-100 was added (1% final concentration) and lysates were incubated for 1 h on ice. The samples were then centrifuged at 4  C for 1 h at 16.000g. Supernatants were stored in aliquots at –70  C. For GST pulldown, equivalent amounts of recombinant proteins (GST or GST–PMM1) were incubated with glutathioneSepharose beads for 1.5 h at 4  C. After washing with lysis buffer 2 (LB2: 10mM Tris–HCl (pH 7.4), 150 mM NaCl, 1 mM MgCl2, 1 mM CaCl2, 0.2% TritonX-100, 5% Trasylol), the loaded beads were incubated with extracts from transfected BHK cells expressing HA–Rab6A Q72R (lane 1), HA–Rab6A T27N (lane 2), or HA–Rab6A wildtype (lane 3). After the beads were washed with LB2, bound proteins were eluted in SDS sample buffer and analyzed by Western blotting.

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Table 5.4 Interaction of Rab6 isoforms with p150Glued hp150Glued 1–1278 214

541

951 1046

735 VK4 523 VK5a

735

Rab6A Q72R Rab6A⬘Q72L Rab6B Q72R

918 ++

+++

++

−

n.t.

−

Results of yeast two hybrid cotransformation assays are shown using Rab6A, Rab6A0 and Rab6B, and p150Glued deletion mutants VK4 and VK5a as constructs. b-galactosidase reporter gene activity was tested.  = no ß-galactosidase activity; ++ = strong b-galactosidase activity; +++ = very high b-galactosidase = MTB (microtubule binding domain); = coiled-coil domain. activity; n.t. = not tested;

In the final part of this review, we show various examples from our coexpression analyses of Rab6 and different interacting partners, including the adaptor protein Mint3 (Figs. 5.8–5.10A and B; Table 5.4). The family of Mint proteins (also called X11 proteins) consists of three members, namely Mint1, Mint2, and Mint3, which all contain a phosphotyrosine-binding domain (PTB) and two carboxyterminal postsynaptic density 95 (PSD-95)/ Discs large/zona occludens-1 (PDZ) domains (Okamoto and Su¨dhof, 1998). All three isoforms interact with the amyloid precursor protein (APP) via their PTB domain (Borg et al., 1996; King and Turner, 2004; McLoughlin and Miller, 1996; Tanahashi and Tabira, 1999). The transmembranal type I protein APP is processed by a, b, and g secretases (Selkoe, 2001). Besides other proteolytic fragments amyloid b-peptide (Ab) is formed, which has been implicated in the pathogenesis of Alzheimer’s disease (Selkoe, 2001). After the detection of Mint3 and Giantin as Rab6 interacting proteins through a YTH screen using a human placenta cDNA library and the GTPase-deficient Rab6A Q72R, colocalization studies were performed (Rosing et al., 2007; Teber et al., 2005). In both cases, a clear partial colocalization signal is present for Rab6 and Mint3 or Rab6A and Giantin, respectively (Figs. 5.8 and 5.9). An alternative approach is shown in Fig. 5.10A and B, where we investigated the interaction of Rab6 and Phosphomannomutase 1 (PMM1). Phosphomannomutases (PMMs) are crucial for the glycosylation of glycoproteins. In humans, two highly conserved PMMs exist, the PMM1 and PMM2 proteins, which in vitro are able to convert mannose-6-phosphate (mannose-6-P) into mannose-1-P. However, only mutations causing a deficiency in PMM2 cause hypoglycosylation, leading to the most frequent type of the congenital disorders of glycosylation (CDG), namely CDG-Ia (Matthijs et al., 1997). PMM1 is as yet

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not associated with any disease, and its physiological role is still unclear. Here, we first tested the interaction of Rab6 with PMM1 applying the YTH system (Fig. 5.10A). Then we confirmed the interaction by a GST pulldown analysis with PMM1 and Rab6 protein (Fig. 5.10B).

4. Concluding Remarks The Rab family’s role in membrane transport specificity is only beginning to be explored and understood. The last years have brought the identification and isolation of numerous effectors of Rab GTPases. Now it is time to characterize their function in more detail and to identify and analyze additional Rab regulator proteins to improve our understanding how the Rab GTPases play their role in intracellular transport processes.

ACKNOWLEDGMENTS We apologize to the authors of the hundreds of papers on small GTPases and interacting proteins that we would have cited had space regulations permitted. The authors would like to thank Bruno Goud, Fumiko Nagano, Casper Hoogenraad, and Anna Akmanova for collaborating on Rab6A/Mint3, Rab6A/p150Glued, Rab6A/BICD. We greatly acknowledge our co-workers M. Bayer, K. Bilbilis, J. Boettcher, E. Bobbert, J. Fischer, C. Holz, M. Kail, M. Kaufmann, J. Kremerskothen, M. Koester, M. Eschricht, T. Matanis, E. Ossendorf, S. Poll-Wolbeck, L. Ruether, M. Rosing, I. Teber, Th. Weide, for contributing to the results summarized in this manuscript. This work was supported by the DFG and an FCI grant to A.B.

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C H A P T E R

S I X

Cubic Membranes: The Missing Dimension of Cell Membrane Organization Zakaria A. Almsherqi,* Tomas Landh,† Sepp D. Kohlwein,‡ and Yuru Deng* Contents 1. Introduction 2. Cell Membrane Architecture 2.1. Membrane symmetries 2.2. Membrane polymorphisms 2.3. Cubic membranes versus cubic phases 2.4. Understanding membrane morphology by transmission electron microscopy 3. Cubic Membranes in Nature 3.1. Cubic membranes: From protozoa to mammals 3.2. Organelles with cubic membrane structure 4. Biogenesis of Cubic Membranes 4.1. Role of membrane-resident proteins in cubic membrane formation 4.2. Role of lipids in cubic membrane formation 4.3. Electrostatic effects on cubic membrane organization 5. Cubic Membranes: Indicators of Cellular Stress and Disease? 5.1. Virus-infected cells 5.2. Neoplasia 5.3. Muscular dystrophy 5.4. Autoimmune disease 6. Cubic Membranes: Specific Functions or Innocent Bystanders? 6.1. Cell space organization and subvolume regulation 6.2. Inter- and intracellular trafficking 6.3. Specific structure-function relationships

* { {

276 278 278 302 303 304 306 306 307 312 313 316 318 319 319 320 320 321 321 322 322 323

Department of Physiology, Yong Loo Lin School of Medicine, National University of Singapore, 117597 Singapore Novo Nordisk A/S, DK-2760 Ma˚lv, Denmark Institute of Molecular Biosciences, University of Graz, A8010 Graz, Austria

International Review of Cell and Molecular Biology, Volume 274 ISSN 1937-6448, DOI: 10.1016/S1937-6448(08)02006-6

#

2009 Elsevier Inc. All rights reserved.

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7. Applications of Cubic Membranes 7.1. DNA transfection 7.2. Do cubic membranes have optical properties? 8. Concluding Remarks Acknowledgments References

324 324 325 326 326 327

Abstract Biological membranes are among the most fascinating assemblies of biomolecules: a bilayer less than 10 nm thick, composed of rather small lipid molecules that are held together simply by noncovalent forces, defines the cell and discriminates between ‘‘inside’’ and ‘‘outside’’, survival, and death. Intracellular compartmentalization—governed by biomembranes as well—is a characteristic feature of eukaryotic cells, which allows them to fulfill multiple and highly specialized anabolic and catabolic functions in strictly controlled environments. Although cellular membranes are generally visualized as flat sheets or closely folded isolated objects, multiple observations also demonstrate that membranes may fold into ‘‘unusual’’, highly organized structures with 2D or 3D periodicity. The obvious correlation of highly convoluted membrane organizations with pathological cellular states, for example, as a consequence of viral infection, deserves close consideration. However, knowledge about formation and function of these highly organized 3D periodic membrane structures is scarce, primarily due to the lack of appropriate techniques for their analysis in vivo. Currently, the only direct way to characterize cellular membrane architecture is by transmission electron microscopy (TEM). However, deciphering the spatial architecture solely based on two-dimensionally projected TEM images is a challenging task and prone to artifacts. In this review, we will provide an update on the current progress in identifying and analyzing 3D membrane architectures in biological systems, with a special focus on membranes with cubic symmetry, and their potential role in physiological and pathophysiological conditions. Proteomics and lipidomics approaches in defined experimental cell systems may prove instrumental to understand formation and function of 3D membrane morphologies.

1. Introduction Membrane-bound cell organelles are typically considered to have rather spherical topology, delineated by one phospholipid-bilayer membrane that separates the interior from the exterior. However, this simplification of organelle topology is a rule not a law, and it is well known that a large number of membrane structures exists in Nature with more complex 3D morphologies. Indeed, the topology of membrane-bound organelles is a rather unexplored area of research. This might be due to

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difficulties in obtaining information about topological parameters from living or fixed cells, and the interpretation of these parameters in the cellular context. Nevertheless, the importance of topology considerations, for example, subcellular compartmentalization, transport phenomena, and sorting events that involve membrane trafficking processes is evident. Cell membrane morphology, controlled by the principles of self-assembly and/or self-organization, is likely to adopt an optimally organized structure under the influence of selective conditions. This is a dynamic process, perhaps restricted to sub-membrane domains, and short-lived, and is dependent on the lipid as well as protein components of the membrane. As a consequence of limited in vivo technologies, knowledge about the molecular mechanisms underlying membrane morphology is scarce and largely restricted to the descriptive level. Indeed, higher order membrane topologies identified by transmission electron microscopy (TEM), are frequently reported in the literature, yet due to their very heterogeneous representations, common features are difficult to comprehend. Among these nonlamellar cell membranes, especially cubic membrane organizations attract great attention (Almsherqi et al., 2006; Hyde et al., 1996; Landh, 1995, 1996) because of their unique feature of 3D periodicity in TEM micrographs and great similarity to the bicontinuous lipidic cubic phases (Bouligand, 1990; Larsson, 1989; Larsson et al., 1980; Luzzati, 1997). Cubic membranes (Figs. 6.1 and 6.2) have therefore often been compared to selfassembled cubic lipidic phases in aqueous dispersions that are well

Figure 6.1 Cubic membrane architecture (Almsherqi et al., 2008). (A) Two-dimensional transmission electron micrograph of a mitochondrion of 10 days starved amoeba Chaos cells and (B) three-dimensional mathematical model of the same type of cubic membrane organization. Scale bar: 250 nm.

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Figure 6.2 Periodic cubic surfaces and cubic membrane. Oblique views of the unit cell of (A) Primitive, (B) Double Diamond, and (C) Gyroid cubic surfaces observed in biological systems. (D) The bilayer constellation of a 3D mathematical model of a cubic membrane. Three parallel Gyroid-based surfaces can be used to describe a biological membrane (bilayer), in which case the centered surface is the ‘‘imaginary’’ hydrophobic mid-bilayer surface and the two parallel surfaces are the two apolar/polar (interfacial) surfaces.

characterized in vitro, with several applications. Indeed, the efforts toward understanding formation and functional roles of cubic membranes in biological systems have been paralleled by the efforts in investigating cubic phases formation and their behavior in lipid–water systems.

2. Cell Membrane Architecture 2.1. Membrane symmetries Biological membranes may exhibit point or line symmetry. A membrane is symmetrical if it can be nontrivially rotated, inverted, mirrored, and translated such that it is indistinguishable from its initial appearance. Symmetry of

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biological membranes is mainly described by rotations. Several sets of membrane arrangements exhibit symmetry such as parallel membranes and hexagonal packing of tubes. In contrast, a cubic membrane exhibits distinct morphological patterns when projected which may even be translated into unique signatures in many directions (Fig. 6.3). The patterned membrane organization of cubic membranes consists of a network arranged in a nonrandom order and is evenly spaced. Therefore, through an overall inspection of TEM micrographs, cubic membranes are recognized via perceptual cues of the patterned membrane organization (Almsherqi et al., 2006; Landh, 1996). This unique appearance of cubic membranes in TEM micrographs frequently allows for the differentiation of cubic membrane organization from other membrane arrangements such as tubulo-reticular structures (TRS) and annulate lamellae (AL) (Figs. 6.4 and 6.5). Cubic membranes represent highly curved, 3D periodic structures that correspond to mathematically well-defined triply periodic minimal surfaces or the corresponding periodic nodal surfaces and their respective constant mean curvature or level surfaces (Fig. 6.2). Both the latter surface descriptions are approximative descriptions of surfaces parallel to the minimal or nonzero level surface (Landh, 1996). Cubic membranes have been detected

Figure 6.3 Computer simulation of TEM images. (A) Schematic illustration of TEM data in 2D projections of a specimen with a finite thickness. A 3D object (a) is depicted and is translucent to the projection rays of an electron beam; (b) representation of one unit cell of the gyroid surface; (c) projection plane onto which the rays impinge, in analogy of the film on which the image would be recorded; (d) 2D projection map provides a corresponding template for matching the patterned membrane domain in the TEM micrograph. (B) Comparison between a 3D cubic membrane model of a gyroidbased surface and its computer simulated projections at different viewing directions. Multiple 2D projections that are generated from the same 3D structure form a library of different patterns. The bottom row corresponds to computer-simulated projections for the top row, based on a projected specimen thickness of one-half of a unit cell viewed at the [1, 0, 0] (left), [1, 1, 0] (middle), and [1, 1, 1] (right) directions. The computergenerated projection can be matched with TEM micrographs to determine the 3D structure of a cubic membrane arrangement (see section 2.4. for further details).

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Figure 6.4 Cell membrane organization. Schematic diagram depicting the likely 3D structure of annulated lamellae, tubulo-reticular structure (TRS) and the membrane folding transition. The pores of annulated lamellae may alternate in arrangement with the symmetry often being quadratic (A) or the pore face each other with the symmetry being hexagonal (B). Two examples of TRS membrane arrangements; (C) interconnected sacular (cisternae) and (D) tubular membrane organization show no global symmetry. A possible model of continuous membrane folding for the formation of double diamond (lower left) and gyroid (upper left) cubic type, hexagonal (upper right) and lamellar structures, and whorls (lower right) (E). The coexistence of these membrane organizations has been reported frequently in UT-1 and COS-7/ CV-1 cells with HMG-CoA reductase and cytochrome b(5) overexpression, respectively. Panels A-D adapted from Figs. 17 and 18; Bouligand, 1991.

without any obvious restrictions or preferences in all kingdoms of life, both under physiological or pathological conditions (Table 6.1). They appear not to be limited to certain types of cells, although they may occur more frequently in some cell types. Furthermore, cubic membranes are not strictly associated with any particular organelle and can apparently evolve from almost any cytomembrane: plasma membrane, endoplasmic reticulum (ER), nuclear envelope (NE, both inner (INE) and outer (ONE)), inner mitochondrial membrane, and the Golgi complex. The smooth ER, however, seems to be the organelle most frequently associated with cubic membrane formation. So far, three surface families have been identified to exist, and these three types of cubic membranes are schematically shown in Fig. 6.2. They are designated according to their corresponding triply periodic minimal (or level) surfaces as gyroid (G), double diamond (D), and primitive (P) surfaces. Cubic membranes often coexist with other ‘‘unusual’’ membrane arrangements, such as TRS, which are irregularly arranged tubes that bifurcate and reanastomose. In many cases, these tubes show a preferential

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A

B

0.5 µm

0.2 µm

C

D

0.2 µm

0.5 µm

Figure 6.5 Examples of different membrane organizations observed in UT-1 cells, 48–72 h after compactin (40 mM) treatment (Deng et al., unpublished). (A) Annulate lamellae, (B) stacked undulated lamellae that show hexagonal transition, (C) cubic, and (D) hexagonal membrane morphologies may coexist in the same cell. Membrane folding appears to originate at the nuclear envelope or the endoplasmic reticulum.

orientation. The main difference of TRS to cubic membranes is that TRS symmetry is usually nematic, since the layers do not show obvious periodic distribution (Fig. 6.4D). The preferential alignment along a direction may be due to an elongation process, perhaps in association with the cytoskeleton, and is not necessarily the result of a spontaneous membrane alignment. However, there are many cases in which the periphery of a perfectly preserved cubic morphology shows TRS appearance, which, therefore, may be introduced by the fixation method (Landh, 1995). Membranes of true cubic morphology are often mis-labeled as TRS in the literature, due to the convoluted image projections observed for both structures. Many of the examples listed in Table 6.1 have been designated as TRS, despite the presence of a distinct cubic symmetry (Almsherqi et al., 2005; Landh, 1996). TRS have attracted biomedical interest due to their potential use as an ultrastructural marker for pathological conditions: they occur in virusinfected and in cancer cells and have been, therefore, often regarded as an

282

Table 6.1 Occurrence of cubic membranes in biological systems. Description of cells/tissue

Monera Gracilicutes Oxyphotobacteria Cyanobacteria Thylakoid lamellae in Anabaena sp. Thylakoid lamellae in Anabaena sp. Thylakoid lamellae in Heterocyst of Anabaena azollae Thylakoid in Anabaena variablis infected with cyanophages Protista Algae Clorophyta Chlorophyceae Membranes in chloroplasts of Zygnema Membranes in chloroplasts of Zygnema Thylakoid membranes in chloroplast of C-10 mutant of Chorella vulgaris Thylakoid membranes in chloroplast of Protosiphon botyoides Charophyceae Plasma membrane of Chara coralline, C. braunii

Sn/a (nm)

References

D/50

Lang and Rae (1967) Beams and Kessel (1977) Lang (1965)

PLB-like structure

P/300

Granhall and von Hofsten (1969)

Quasi-crystalline lamellar

Pm/350 Gm/500

McLean and Pessoney (1970) Deng and Landh (1995) Bryan et al. (1967)

Cognomes

Honeycombed lamellae

Masses of prethylakoid tubules Sinusoidal thylakoids

Charasome

Berkaloff (1967)

G/140

Barton (1965), Franceshi and Lucas (1980, 1981), Lucas and Franceshi (1981)

Plasma membranes of Nitella Rhodophyta Rhodophyceae ER in Erythrocystis montagnei Gymnomycota (Myxomycota, slime moulds) Plasmodiogymnomycotina Myxomycetes Mitochondria in Physarum polycephalum Mitochondria in Didymium nigripies Mastigomycotina Diplomastigomycotina Oomycxetes ER in Oedogoniomyces zoospores

Interconnected tubules

G

Crystalline body

Crawley (1965)

Tripodi and de Masi (1977)

Regular tubular network Unusual tubular morphology

D2 D2

Daniel and Ja¨rlfors (1972a,b) Schuster (1965)

Organized lamellar system

G/215

Reichle (1972)

Protozoa Sarcomastigophora Mastigophora Phytomastigophora Photosynthetic lamellae in dark-grown Meshed network, Chlamydomonas reinhardi y-1 PLB-like Zoomastigophorea ER in Leptomonas collosoma Membrane lattice Rhizopodea Amoebida Mitochondria in C. carolinense

Friedberg et al. (1971) Hoober et al. (1969)

283

D/88

Linder and Staehelin (1980)

D2/150

Pappas and Brandt (1959), Brandt and Pappas (1959), Borysko and Roslansky (1959), Daniels and Breyer (1968) (continued)

284

Table 6.1 (continued) Description of cells/tissue

Cognomes

Sn/a (nm)

References

Complex tubular patterns

D2, P2/ 130 D2

Deng and Mieczkowski (1998); Deng et al. (1999) Daniels and Breyer (1965)

Mitochondria in C. carolinensea Mitochondria in Chaos illinoisensis Pelobiontida Intranuclear membrane in Pelomyxa palustris Cnidospora Microsporidea Membranes in sporoblast of Nosema apis Ciliophora Oligohymenophora Hymenostomatida Pniculina Contractile vacuolar membranes of Paramecium aurelia, Paramecium multimicronucleatum

Intranuclear membrane of Neobursaridium gigas Tetrahymenina Contractile vaculor membranes of Tetrahymena pyriformis

Fungi Amastigomycota Ascomycotima

Crystalloid

Daniels and Breyer (1967)

Honeycomb network

Youssef and Hammond (1971)

Smooth spongiome

Mckanna (1976), Allen and Fok (1988), Allen et al. (1990), Fok et al. (1995), Hausmann and Allen (1977), Ishida et al. (1993, 1996) Nilsson (1969)

Crystal configuration

Nephiridial tubules

G/160

Elliott and Bak (1964)

Ascomycetes ER in apothelial cells of Ascobolus stercorarius Plant Pteridophyta Oocytes in Selaginella kraussiana Sprematohyta Angiosperms Magnoliophyta (anthophyta) Dicotyledons Ranunculidae Ranunculaceae ER of nectaries in Helleborus foetidus

285

ER of ovules in Ficaria ranunculoides ER in virus-infected leaf parenchyma cell of Helleborus niger ER in phloem-parenchyma cells of Helleborus lividus ER in differentiating sieve elements of Eranthis cilicica Papaveraceae ER in ovules of Papaver rhoeas Hamamelididae Urticales ER in differentiating sieve elements of Ulmus americana

Lattice bodies

G/55

Pseudocrystal

Anderson and Zachariah (1972), Zachariah (1970), Zachariah and Anderson (1973), Wells (1972)

Robert (1969a,b)

Cotte de mailles

P2/80

Cotte de mailles ER complex

P P2/65

Eyme´ (1963, 1966), Eyme´ and Blance Le (1963) Eyme´ (1966, 1967) Robinson (1985)

ER complex

P2/75

Behnke (1981)

ER complex

G2/70, 145

Behnke (1981)

Cytoplasmic complex

P2, D2

Ponzi et al. (1978)

Complex network/maize

Evert and Deshpande (1969)

(continued)

Table 6.1

(continued)

286

Description of cells/tissue

Caryophyllidae Caryophyllales ER in sieve elements of beet yellow vein virus infected Beta Dilleniidae Capparales ER in nectaries of Diplotaxia erocoides Malvales ER in differentiating sieve elements of Gossypium hirsutum Rosidae Leguminosae Plastids in bean root tips of Phaseolus vulgaris ER in differentiating sieve element of P. vulgaris Sapindales ER in differentiation sieve elements of Acer ER in differentiation sieve elements of Acer pseudoplatanus Asdteridae Gentianales ER in differentiating sieve element of Nymphoides peltata Monocotyledons Commelinidae Poales

Cognomes

Sn/a (nm)

Convulated ER

Cotte de mailles

References

Esau and Hoefert (1980)

P2/55

Eyme´ (1966, 1967)

Convoluted ER

Thorsch and Esau (1981)

Tubular complex

Newcomb (1967)

Convoluted membranes

Esau and Gill (1971)

Quasi-crystalline membranes Vesicular aggregates

D2/180

Convoluted membrane complex

G2/125

Wooding (1967) Northcote and Wooding (1966)

Johnson (1969), Oparka and Johnson (1978)

Poacea (Gramineae) ER in Triticum aestivum infected by Membranous body wheat spindle streak mosaic virus

287

Liliidae Liliales ER of differentiating sieve elements Dioscorea bulbifera ER of differentiating sieve elements Dioscorea macroura ER of differentiating sieve elements Dioscorea reticulata ER of differentiating sieve elements Smilax excelsa Arecidae Arecales ER in differentiating sieve elements of Cocus nucifera ER in differentiating sieve elements of Chamaedorea pulchra, C. oblongata, C. elegens, Elaeis guineensis Gymnosperms Coniferophyta Conniferales ER in sieve cells in Pinus strobus ER in sieve cells in Pinus pinea Animalia Mollusca Cephalopoda Nautiloidea

G2

Hooper and Wiese (1972), Langenberg and Schroeder (1973)

Lattice-like membrane

G/40

Behnke (1968)

Lattice-like membrane

G1, G2/ 30, 140 G/35

Behnke (1968)

Lattice-like body Convoluted ER

Convoluted tubular ER

Behnke (1965, 1968) Behnke (1973)

G2

Parthasarathy (1974a,b)

Convoluted tubular ER

Parthasarathy (1974a,b)

Lattice-like bodies Vesicular aggregation

Murmains and Evert (1966) Wooding (1966)

(continued)

Table 6.1

(continued)

288

Description of cells/tissue

Nautilida ER in retinula cells in Nautilus macromphalus Gastropoda Opisthobranchia Nudibranchia ER in spermatids of Spurilla nepolitana

Pulmonata Helicidae ER in photoreceptor cells of Helix pomatia Basommatophora ER in spermatids of Planorbarius corneus Stylommatophora ER in type I photoreceptor cells of Limax maximus Annelida Polychaeta Aphroditidae, Polynoı¨nae ER in luminous cells of Acholoe astericola

ER in luminous cells of Lagisca extenuata ER of photoreceptor cells in L. extenuata

Cognomes

Sn/a (nm)

References

Tubular array of myeloid body

P > 10/ 550

Barber (1967), Barber and Wright (1969)

Undulating tubular body

P2/130

Eckelbarger (1982), Eckelbarger and Eyster (1981)

Biocrystal

Ro¨hlich and To¨ro¨k (1963)

Cytoplasmic crystalloid

D2/50

Starke and Nolte (1970)

Corrugated ER

D2/195

Eakin and Brandenburger (1975)

PER, Photosomes

D2/250

PER PER

D2/250 D2

Bassot (1964, 1966), Bassot and Nicolas (1978), Bilbaut (1980), de Ceccatty et al. (1977) Bassot (1966) Bassot and Nicolas (1978)

ER in luminous cells of Harmothoe lunulata ER of photoreceptor cells in Arctonoe vittata Syllidae ER of photoreceptor cells in Syllis amica Nereidae ER of inner segment in photoreceptor cells in Nereis virens ER of photoreceptor cells in Nereis limnicola Oligichaeta Lumbricidae ER in spermatids of Eisenia foetida Hirudinea Gnathobdeliae ER in photoreceptor cells of Hirudo medicinalis Arthropoda Arachnida Scorpions Mitochondria in sprematids of Euscorpius flavicaudis Pseudoscorpions ER of spermatids in Diplotemus sp.

PER

Bassot (1985), Bassot and Nicolas (1987, 1995), Nicolas (1979, 1991) Singla (1975)

Crystalline element

PER

D2/50

Paracrystalline body Crystalloid body

Bocquet and DhainautCourtois (1973) Dorsett and Hyde (1968)

G

Eakin and Brandenburger (1985)

Undulating tubular body

Stang-Voss (1972)

PER

Walz (1982)

Andre´ (1959)

Highly ordered membrane

Bawa and Werner (1988)

289

(continued)

Table 6.1

(continued)

290

Description of cells/tissue

Crustacea Copepoda ER of retinula cell in Macrocyclops albidus Malacostraca Decapoda ER of spermatozoa in Cragon septemspinosa Schwann cell processes in the ventral nerve cord of Procambarus sp. Schwann cell processes in the walking limb nerves Nephrops sp. Mitochondria in oocytes of Cambarus and Orconectes Isopoda ER in bordering cells of Bellonci organ in Sphaeroma serratum Tanaidacea ER in sperm of Tanais cavolinii Insecta Apterygota Thysanura ER in rectal epithelial cells of Petrobius maritmus Mitochondria in intestinal cells of P. maritmus Pterygota Orthoptera ER in spermatids of Melanoplus diffentialis differentialis

Cognomes

Sn/a (nm)

Elaborately wound membranes

Paracrystalline lattice

References

Fahrenbach (1964)

D2

Arsenault et al. (1979, 1980)

Anastomosing tubular inclusion Anastomosing networks

Pappas et al. (1971)

Honeycombed cristae

Beams and Kessel (1963)

Annulate lamellae

Holtzman et al. (1970)

G/50

Spongy/foamy cytoplasm

Puzzles tridimensionnels

Cotelli and Donin (1980)

G2/120 D2/160

Textum

Chaigneau (1971)

P2/250

Fain-Maurel and Cassier (1972) Fain-Maurel and Cassier (1973)

Tahmisian and Devine (1961)

Mitochondria in corpus allata of Locust migratoria migratorioides Hemiptera ER in spermatids of Dysdercus fasciatus ER in oocytes of Pyrrocoris apterus ER in spermatogenic cells of Notonecta undulata ER in spermatogonai a and spermatocytes of P. apterus Diptera Mitochondria in flight muscle cells of Calliphora erythrocephala ER in photoreceptor cells of vitamin A deficient Aedes aegypti Lepidoptera SER in scale cells of butterfly Mitoura grynea Hymenoptera ER in secretory cells of Dufour’s gland in Parischnogaster mellyi Blattodea Mitochondrion in secretory cells of the spermatheca in Periplaneta am. Chordata Urochordata Ascidiacea tethyodea Stolidobranchiata

Fain-Maurel and Cassier (1969) Sinusoidal tubules PER Anastomosing tubules

D2/150 D2/250

Folliot and Maillet (1965) Mays (1967) Tandler and Moribier (1974)

PER

G2/175

Wolf and Motzko (1995)

Regular fenestrated cristae Masses of membranes

Smith (1963)

Membrane-cuticle unit

Chiradella (1989, 1994)

Vesicular profiles

Delfino et al. (1988)

Brammer and White (1969)

Gupta and Smith (1969)

291

(continued)

292

Table 6.1

(continued)

Description of cells/tissue

Golgi of test cells in the ovary of Styela sp. Cephalchordata ER in Joseph’s cells of the Branchiostoma lanceolatum Vertebrata Agnatha; Cephalaspidomorphii Petromyzoniformes ER in retinal pigment epithelium cells of Lampetra fluviatalis Osteichthyes Actinopterygii Salmoniformes ER in epithelium of the olfactory organ in Salmo trutta trutta Plasma membrane in gill epithelia cells of Salmo salar ER in adrenocortical cells of Salmo fario Siluriformes ER of clear cells in the dendritic organ of Plotsus anguillaris Anguilliformes ER in ‘‘club cells’’ of juvenile Anguilla rostrata

Cognomes

Sn/a (nm)

Honeycomb, lattice-like

References

Kessel and Beams (1965)

Meandrous tubules

G2/175

Welsch (1968)

Undulated membrane complex

G4/155

¨ hman (1974) O

Turtuous interconnected ER Tubular system

Bertmar (1972) D

Pisam et al. (1995)

Imbricated cisternae of ER

G2/200

Jung et al. (1981)

Tubular network

D/100

van Lennep and Lanzing (1967)

Array of circular figures

Leonard and Summers (1976)

293

Perciformes ER in chloride cell of freshwateradapted Scophthalmus maximus Plasma membrane in gill epithelia cells of Oreochromis niloticus Dipnoi Lepidosireniformes ER of Neuroepithelial cell in the lung of Protopterus aethiopicus Crossopterygii ER in retinal pigment epithelium cells of Latimeria chalumnae Amphibia Anura Pipidae Mitochondria in Sertoli cells of Xenopus laevis Discglossidae ER in intestinal epithelium cells of Alytes obstetricans Ranidae ER in secretory gland of Dendrobatidae anthony, D. auratus Urodel Salamandridae ER in oocyte of Necturus maculosus maculosus ER in retinal pigment epithelium cells of Notophtalamus viridescens

Membraneous tubular system Tubular system

D

Pisam et al. (1990)

D

Pisam et al. (1995)

Adriaensen et al. (1990)

Paracrystalline inclusion

Regular arrays of tubules

G

Locket (1973)

Regularly fenestrated cristae

D2/105

Kalt (1974)

Sinusoidal tubules

Crystalloid

Hourdry (1969)

G

Neuwirth et al. (1979)

Annulate lamellae

Kessel (1990)

Fenestrated lamellae

Yorke and Dickson (1985)

(continued)

Table 6.1

(continued)

294

Description of cells/tissue

Bufonidae ER of cells in the parotoid gland of Bufo alvarius ER in spermatids of Bufo arenarum Reptilia Lepidosauria Squamta ER in spermatids in Anolis carolinensis Aves Galliformes ER in retinal pigment epithelia cells of Cortunix cortunix japonica ER of epithelium in uropygial gland of Cortunix cortunix japonica Mammalia Scandentia Tupaiidae Mitochondrias in photoreceptor cone cell of Tupaia glis SER of cells in the adrenal cortex T. glis Mitochondria in retinal cone cell of Tupaia belangeri Chiroptera Molossidae ER of cells in sebaceous gland of Tadarida brasiliensis

Cognomes

Sn/a (nm)

References

Crystalloid

Cannon and Hostetler (1976)

Annulate lamellae

Cavicchia and Moviglia (1982)

Membranous body

Clark (1967)

Ahn (1971) Crystaloid

Fringes and Gorgas (1993)

Concentric whorls of cristae Crystalloid

G10/500

Samorajski et al. (1966)

D

Hostetler et al. (1976)

Peculiar whorls of cristae

G12/400

Foelix et al. (1987), Knabe and Kuhn (1996), Knabe et al. (1997)

Crystalloid

D2/105

Gutierrez and Aoki (1973)

Carnivora Felidae ER of bright columnar cells in the vomeronasal organ of the cat Canidae ER of follicular cells in adenohypophysis of the dog ER in cutaneous histiocytoma cells of the dog ER in adventitial cells of the dog ER in mononuclear cells of dog treated with anti-dog-lymphocyte serum Lagomorpha Leproidae ER in ovarian steroid cells of the rabbit ER of type II cells in taste buds of male albino rabbit ER in endothelial cells and macrophage of the New Zealand white rabbit infected with herpes simplex virus Ochotonidae ER in Mu¨ller cell of Ochotona sp.

Hexagonal crystal-like membrane

G

Seifert (1971, 1972, 1973)

(Tweedlike) paracrystal

Nunez and Gershon (1981)

Paracrystal

Marchal et al. (1995)

Tubular aggregates Inclusion body surrounded by limiting membrane

Blinzinger et al. (1972) Somogyi et al. (1971)

Blanchette (1966a, b) Toyoshima and Tandler (1987) Baringer and Griffith (1970)

Crystalline aggregates

Well-developed networks of ER

295

Artiodactyla Suidae ER in skin cells of pig infected with swine Cytoplasmic inclusion pox virus

G2/315

Hirosawa (1992)

Cheville (1966)

(continued)

Table 6.1

(continued)

296

Description of cells/tissue

ER in endothelial cells of cervical cord of the pig infected with virus Bovidae ER of cell in preputial gland of female Capricornus cripus Intranuclear tubules in bovine tissue with papulosa-virus infection Perissodactyla Equidae ER in sebaceous gland of Equidae Rodentia Muridae ER in rat renal tubule cells ER in rat hepatocytes after hexachlorohexahydrophenanthrene in diet ER in hepatocytes of carbon tetrachloride fed rats ER in rat hepatocytes after phenobarbital treatment ER in hepatomas of the rat ER in lutein cells of the rat after cycloheximide treatment ER in adrenal medullary cell of chlophentermine treated rat ER in adrenal cortical cell of chlophentermine treated rat ER in meibomian glands of the rat Mitochondria in skeletal muscle of the rat

Cognomes

Sn/a (nm)

Koestner et al. (1966)

Crystal arrays

Grids of SER

G/80

Intranuclear tubule-like structure

Grids of SER

References

Atoji et al. (1989) Pospischil and Bachmann (1980)

G

Jenkinson et al. (1985)

Fenestrated membranes Flattened vesicles

Bergeron and Thie´ry (1981) Norback and Allen (1969)

Labyrinth tubular aggregates Meshed network

Reynolds and Ree (1971)

Crystalline tubular aggregates Crystalloid body Dense body

Bolender and Weibel (1973) Hruban et al. (1972) Horvath et al. (1973) Lu¨llmann-Rauch and Reil (1973) Lu¨llmann-Rauch and Reil (1973) Sisson and Fahrenbach (1967) Leeson and Leeson (1969)

ER in jejunal absorptive cells of rat intestine ER in vomeronasal epithelium in the rat ER in cell of sebaceous gland in mouse skin ER of neurons in the mice ER in testicular interstitial cells of mice ER in Leydig cells of mice ER in mice retinal pigment epithelium after mild thermal exposure ER in hepatocytes of chlophentermine treated mice ER in hepatocytes of mice infected with mouse hepatitis virus ER in mice brain cells inoculated with St. Louis encephalitis virus ER in neuron of suckling mouse infected with Semliki Forest virus Cricetidae ER in UT-1 cells with HMG-CoA reductase expression ER of CHO cells with rubella virus E1 glycoprotein expression ER in hepatocytes of the hamster after phenobarbitone treatment ER in sebaceous gland of the hamster

Hatae (1990) Membranous body Crowded elements

Garrosa and Coca (1991) Rowden (1968)

Interconnected segments of SER Network of tubules

Johnson et al. (1975) Christensen and Fawcett (1966) Russel and Burguet (1977) Kuwabara (1979)

Tubular profiles Lacy patterened ER Crystalline-like body

Lu¨llmann-Rauch and Reil (1973) Ruebner et al. (1967)

Peculiar tubular structures Convoluted membranous mass Anastomosing membrane tubules Sinusoidal ER

Murphy et al. (1968) Grimley and Demsey (1980, p. 151) D2/245

Pathak et al. (1986)

Tubular membrane

Hobman et al. (1992)

Membrane complex

Ghadially (1988, p. 512)

Grid of SER

Bell (1974a)

297

(continued)

Table 6.1

(continued)

298

Sn/a (nm)

References

Description of cells/tissue

Cognomes

ER in sebaceous gland of androgen treated hamster ER in smooth muscle cell of triparanol treated male hamster Mitochondria in serous secretory cells of Meriones unguiculates Subdermal tumour in the hamster produced by inoculation of: M-1 Caviidae ER in adrenal cortical cells of fetal guinea pig, Cavia sp. ER in receptor cells in the vomeronasal organ of newborn Cavia sp. Primates Strepsirhini (Prosimii) ER in sebaceous gland of Galago senegalensis ER of interstitial cells in the antebrachial organ of Lemur catta Haplorihini Tarsiiformes ER in sebaceous gland of Tarsier syrichta Simiifomes Cerchopithecidae: ER in CV-1 cells infected with simian virus 40 ER in COS cells upon overexpression of msALDH ER in COS-7 and CV-1 cells upon overexpression of cytochrome b(5)

Grids of SER

Bell (1974b)

Dense bodies

Chen and Yates (1967) P2/175

Undulating tubules, UMS

Spicer et al. (1990) Chandra (1968)

Black (1972) Mendoza and Ku¨hnel (1989)

Tubules of SER

G, P

Bell (1974a)

Crystalloid

G/70

Sission and Fahrenbach (1967)

Grids of SER

D2

Bell (1974a) Kartenbeck et al. (1989)

Tubular membranes network Crystalloid Organized SER

Yamamoto et al. (1996) G2, D2

Snapp et al. (2003)

299

ER in Vero cells infected with SARS coronavirus ER in sinusoidal endothelial cell in liver of Macaca fascicularis Mulatta: ER in sebaceous gland of Macaca menstrina ER in retinal epithelium cells of Macaca mulatta ER in endothelial cells of the glomerular capillaries in M. Mulatta ER in endothelial cells in liver of M. mulatta Ibidem with nutritional cirrhosis ER in epidermal pox disease of M. mulatta ER in spinal/endothelia cells of M. mulatta after tumor induced by sarcoma virus ER in kidney cells of M. mulatta infected with Tana poxvirus ER in macrophages, neutrophilic granulocytes and plasma cells of M. mulatta infected with SIV ER in monkey kidney CMK cells infected with poliovirus ER in endothelial cells of monkey spinal cord infected with poliovirus ER in MA 104 cells infected with Simian rotavirus SA11 ER in LLC-MK2 infected with rubella virus

Tubuloreticular structures Crystalloids

G

Goldsmith et al. (2004) Tanuma (1983)

Tubules of SER Peculiar body

Bell (1974a,b) Ishikawa (1963)

Round of hexagonal bodies Cytoplasmic crystalloid

de Martino et al. (1969)

Cytoplasmic crystalloid Crystalloid Crystalline inclusion

Ruebner et al. (1969) Casey et al. (1967) Munroe et al. (1964)

Honeycombed crystals

Espan˜a et al. (1971)

Tubuloreticular structures

Kaup et al. (2005)

Paracrystalline arrays

Hashimoto et al. (1984)

Paracrystalline arrays

Blinzinger et al. (1969)

Smooth membrane vesicles Crystal lattice-like structure

Quan and Doane (1983)

Ruebner et al. (1969)

Kim and Boatman (1967) (continued)

Table 6.1

(continued)

300

Description of cells/tissue

Ceboidea: ER in rous sarcoma virus induced tumour cells of Saquinus sp. Hominoiddea: ER in hepatocytes of d-agent inoculated Pan trodeglytes ER in hepatocytes of P. trodeglytes post experimental hepatitis ER in endothelial cells of human and chimpanzee liver infected with hepatitis virus Man: ER in villus absorptive cells in fetal small intestine of man ER in cells of adrenal gland in man ER in HEp-2 cells infected with Ilheus virus ER in human cancer cell lines: F-3, -9, -24, -53, No. 2117 ER in HeLa cells ER in HT-29 cells infected with rotavirus ER in cells from lymph-node culture of a patient with reticulum-cell sarcoma ER in B lymphocyte of a 6-month-old male infant ER in endothelial KS cells ER in P3-J cells ER in human lymphocytes Mitochondria in adenoma of submandibular gland of man

Cognomes

Membrane inclusion

Sn/a (nm) 2

P /175220

References

Smith and Deinhardt (1968) Canese et al. (1984)

UMS

Pfeifer et al. (1980)

Tubuloreticular and paracrystalline inclusion Convoluted membrane

Schaff et al. (1992)

Knotted membranes UMS

Moxey and Trier (1979) McNutt and Jones (1970) Tandler et al. (1973) Chandra (1968)

Cotte de maillet Tubuloreticular structures Membrane inclusion with crystalline pattern Tubular arrays

Franke and Scheer (1971) Tinari et al. (1996)

Paracrystalline inclusions UMS Microtrabecular lattice Reticulate cristae

Marquart (2005) Chandra and Stefani (1976) Guatelli et al. (1982) Tandler and Erlandson (1983)

Moore and Chandra (1968) Geha et al. (1974)

Mitochondria in metastatic melanoma in man Lysosomes in human myxoid chondrosarcoma ER in human embryonic kidney cells infected with HRV ER in epithelial lung carcinoma of man Inner nuclear membrane in parosteal sarcoma of man ER in bronchiogenic carcinomas ER of endothelial cells on glomerular capillaries of nephritic man ER of endothelial cells in a hepatoblastoma of man ER in soft tissue sarcoma of man

Ghadially (1988, p. 212) Cameron et al. (1992) Ghadially (1988, p. 496)

Vesicular structure Micro- TRS TRS UMS

D

Schaff et al. (1976) Murray et al. (1983)

Crystalline bodies

Ghadially (1988, p. 96) de Martino et al. (1969)

TRS

Gonzales-Crussim and Manz (1972) Szakacs et al. (1991)

Sn/a Indicates that the membrane (surface) morphology (S) is consistent with (n) membranes or multiple (m) membranes with a lattice size of (a). The listed unit cell size is based on either DTC analysis or direct measurements of the 2D lattice parameters. UMS: undulating membrane structure; TRS: tubuloreticular structure; PER: paracrystalline ER; SER: smooth endoplasmic reticulum. The table summarizes the observation of cubic membranes in normal, pathological, and experimentally manipulated cells.

301

302

Zakaria A. Almsherqi et al.

indicator of infection or transformation. For example, TRS have been observed in cells infected with SARS (Almsherqi et al., 2005) and HIV (Kostianovsky et al., 1987). In addition to TRS, annulate lamellae (AL) are another type of convoluted 3D membrane structure, and their appearance is also often correlated with that of cubic membranes. AL are frequently observed in differentiating gamates, namely in vertebrate oocytes and in spermatogonia, and appear to occur in close association with the cell nucleus (see Table 6.1). Tangential TEM sections of AL most often exhibit a hexagonal arrangement (Kessel, 1983), whereas perpendicular sections do not reveal any obvious symmetrical arrangement, even though they always exhibit an astonishingly regular organization, indicating an underlying periodic structure. Based on the apparent morphological similarities between AL and the NE, it has been suggested that AL represent a cytoplasmic NE extension that functions as a reservoir for both ER membrane components and nuclear pores (Kessel, 1983, 1992). In favor of such speculations is the fact that AL have been observed in direct continuity with the outer nuclear membrane, and that they also have been suggested to contain nuclear pore complexes (Landh, 1996). AL are assembled of superimposed pairs of membrane bilayers, which join along the pores whose distribution may vary (hexagonal, quadratic, or random). The pores present in AL are either facing each other if the membrane symmetry is hexagonal (Fig. 6.4B) or the appearance of pores alternates in a quadratic membrane arrangement (Fig. 6.4A).

2.2. Membrane polymorphisms The coexistence of different subtypes of cubic membranes or together with other membrane organizations within the same cell organelle is quite frequent, pointing to structural or functional relationships between these membrane arrangements (Fig. 6.4E). Probably the most evident example is the ER where different membrane morphologies such as cubic membranes, lamellar and hexagonal membranes, and whorls coexist quite commonly (Landh, 1996; Snapp et al., 2003). Coexistence of at least two cubic membrane subtypes within the same organelle has also been observed in mitochondria of amoeba Chaos carolinense (Deng and Mieczkowski, 1998). In this organism, the relative abundance of gyroid (G) or diamond (D) and primitive (P) subtypes of cubic morphology changes during starvation, the biological significance of this polymorphic behavior, however, is currently unknown. The ease with which cubic membranes and other membrane arrangements are interconverted can be attributed, at least in part, to the effect of weakly dimerizing ER proteins (Snapp et al., 2003). Previous work suggested that crystalloid ER biogenesis entailed a tight, zipper-like dimerization of the cytoplasmic domains of certain ER-resident proteins

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(Yamamoto et al., 1996). However, Snapp et al. (2003) found that organized smooth ER (OSER)-inducing proteins can trigger cubic membrane formation upon over-expression through low-affinity interactions between cytoplasmic domains. This observation might explain phenomena such as (a) the heterogeneity of ER membrane structures, (b) the high rate of (reversible) lamellar to cubic membrane transition, and (c) the technical difficulties and limitations in isolating intact cubic membranes from biological samples.

2.3. Cubic membranes versus cubic phases Lipidic bicontinuous cubic phases consist of hyperbolically curved bi-layers where each monolayer is draped over a periodic cubic (minimal) surface (Fig. 6.2D). With respect to bilayer arrangements, the geometries of cubic membranes are similar to those of the cubic phases, however, two major differences exist: (i) the unit cell size and (ii) the water activity. It has been argued that the latter must control the topology of the cubic membrane (Bouligand, 1990) and hence that the cubic membrane structures must be of the inverted type rather than ‘‘normal’’ type (type I). All known lipid–water and lipid–protein–water systems that exhibit phases in equilibrium with excess water are of the inverted type (type II). Thus, water activity alone cannot determine the topology of cubic membranes. Inverted cubic phases have been observed with very high water activity (70–90%), in the mixtures of lipids, in lipid–protein systems, in lipid–polymer systems (Landh, 1994), and in lipid and lipopolysaccharide mixtures (Brandenburg, 1990, 1992). Most cubic phases in lipid–water systems exhibit unit cell parameters not larger than 20 nm, while in cellular cubic membranes the lattice size is usually larger than 50 nm. However, in lipid–protein–water, lipid–poloxamer–water and lipid–cationic surfactant–water systems, cubic phases with cell parameters of the order of 50 nm have also been observed (Landh, 1996). On the other hand, the unit cell size of cubic membranes is rarely less than 50 nm (e.g., in prolamellar bodies) and the size ranges from 50 to 500 nm. Cubic membranes with large lattice size (500 nm) were frequently observed in chloroplast membranes of green algae Zygnema (Fig. 6.6). Additionally, cubic membranes are formed under conditions corresponding to a highly regulated multiphase ‘‘equilibrium’’ process. This is supported by the fact that they are usually formed in close contact with different other membrane configurations. The asymmetry of biological membranes with respect to the two leaflets is likely to affect cubic membrane formation, in particular as a consequence of lipid and protein composition, and interaction with the surrounding ion milieu.

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Figure 6.6 Multilayer membrane organization and transformation. (A) An overview of the ultrastructure of chloroplast membrane in green algae Zygnema sp. (LB923) at 41 days of culture. Scale bar: 1 mm. (B) Several subdomains display different morphologies, ranging from simple stacked lamellar in direct association with paired parallel membranes (2 membranes; upper left) and double paired parallel membranes (4 membranes; lower right) of the gyroid-based cubic membrane morphology. Scale bar: 500 nm (Deng, 1998).

2.4. Understanding membrane morphology by transmission electron microscopy A survey of the literature (Table 6.1) immediately unveils a multitude of ‘‘unusual’’ membrane organizations in various cell types. Most of these depictions were obtained by TEM of chemically fixed and thin-sectioned cells and tissues. Dependent on the thickness and orientation of the section through the specimen, relative to the coordinates of an ordered 3D structure, various types of projection patterns are observed. As a consequence, membrane ultrastructures derived from TEM images are frequently misinterpreted, in particular for the highly folded and interconnected 3D morphologies resembling cubic membranes. TEM relies on 70–90 nm thick sections through the specimens and the 2D image obtained is the result of a projection of a 3D structure. Therefore, nonlamellar biological membranes, such as inverted hexagonal or cubic structures, may yield very heterogeneous projection patterns by TEM, dependent on the orientation of the section relative to the structural axes (Fig. 6.3). Interpretation of TEM membrane patterns is further complicated if the lattice size of the observed structure is considerably smaller than that of the section thickness.

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Serial sections or scanning EM, as well as tilting and rotation of the sample, may facilitate structure interpretation. Furthermore, TEM of multiple randomly cut sections through a specimen provides a rather simple means to reconstruct its 3D structure. More elaborate electron tomography (ET) has contributed a great deal of resolution to understanding cubic membrane organizations and their continuity with and relations to the neighbor structures (Deng et al., 1999). In ET, rather thick sections (400 nm) are imaged in multiple tilted angles (up to
60 ), yielding a large number of projections; these images are reconstructed by computational image analysis into a 3D representation of the object, which allows the 3D reconstruction of cellular structures with a resolution of 5 nm, that is, approaching the level or larger molecular assemblies (for a review see Lucic et al., 2005). EM tomography has previously been successfully applied to determine cubic membrane transition of the inner mitochondrial membrane morphology in the amoeba C. carolinense upon starvation (Deng et al., 1999). Cryo-ET from specimens in vitreous ice further improves sample preservation and membrane resolution, but obviously is not yet routinely established. Cryo-ET avoids common artifacts of conventional EM preparation techniques and is also suited for high-resolution analyses of membrane-bound organelles (Hsieh et al., 2006; Lucic et al., 2005). Most EM experiments described in the literature that focus on biological membranes were obviously not designed to depict three-dimensionally convoluted membrane arrangements. Therefore, alternative methods have to be applied to reconstruct—potential—3D membrane morphologies from single TEM sections. Indeed, based on well-defined mathematical models of cubic membrane arrangements, projections can be calculated that simulate various section orientations and thicknesses (Fig. 6.3). Such a ‘‘direct template correlative’’ (DTC) matching method (Almsherqi et al., 2005, 2006; Deng and Mieczkowski, 1998; Landh, 1995, 1996) has been developed based on pattern and symmetry recognition. Through the DTC method, the electron density of the TEM image is correlated to a library of computer-simulated 2D projection maps that allows to unequivocally deduce the nature of the cubic membrane arrangement. An application of the DTC method to identify cubic membrane organization in TEM micrographs is shown in Fig. 6.7. In brief, the 2D projections (Fig. 6.7C) calculated from a mathematical 3D model (Fig. 6.7B) are matched with a selected TEM micrograph (Fig. 6.7A); consequently, a successful pattern match defines the nature of the membrane arrangement in 3D (Deng and Mieczkowski, 1998; Landh, 1995, 1996). The DTC method simplifies the experimental requirements for recording cubic membranes in biological samples, and can also be applied to examine published TEM micrographs in retrospect. The following section highlights the identification of cubic membrane structures in multiple cellular systems and subcellular organelles.

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Figure 6.7 Direct template matching method. (A) TEM micrograph of lens mitochondria observed in the retinal cones of tree shrew species; (B) 6 pairs (12 layers) of G-based parallel level surfaces—a mathematical 3D model—that can be used to describe G type of cubic membrane morphology and the corresponding computersimulated 2D projection map (C) derived from the corresponding 3D model in (B) (image provided by Prof. S. Wagon, St. Paul, Minnesota); TEM micrograph of lens mitochondria (A) perfectly match the theoretical projection (C), that is generated from 6 pairs (or 12 layers) of G-level surfaces (
0.1,
0.2,
0.4,
0.5,
0.7,
0.8) with a quarter of a unit cell section thickness viewed from the lattice direction [1, 1, 1]. Note the matching details of the TEM projection and computer-simulated 2D projection such as the appearance of density of the lines (membranes) and the density between the sinusoid membranes. The original TEM micrograph in (A) is adopted from Fig. 6.10, from Foelix et al. (1997) with kind permission of Springer Science and Business Media. (14,000 ).

3. Cubic Membranes in Nature 3.1. Cubic membranes: From protozoa to mammals Extensive membrane proliferations leading to unusual and highly convoluted depictions in TEM micrographs have been observed in numerous cell types from all kingdoms of life and in virtually any membrane-bound subcellular organelles, as outlined above. Table 6.1 summarizes a survey of the literature of the past six decades on cubic membrane morphologies identified in organelles, from protozoan to human cells. The occurrence of cubic membranes is listed by genera and, if applicable, the type and lattice size of the cubic membrane extracted from the published

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TEM images are presented (see also Hyde et al., 1996; Landh, 1996). Not surprisingly, due to the absence of a clear understanding of the 3D structure of the depicted membranes, many of the examples have been considered as novelties with little or no reflection on the wealth of related contributions in the literature. Hence, these morphologies appear under a large variety of nicknames, some of which are also listed in Table 6.1. Furthermore, the examples have been chosen to best represent the structural characteristics of cubic membranes, and an effort has been made to leave out those perhaps less recognizable structures such as ‘‘membraneous tubular’’, ‘‘cisternal systems’’, ‘‘tubular inclusions’’, or ‘‘cisternal convolutions’’ etc. In many cases where we have chosen not to classify the cubic membrane it is mainly due to the lack of discernible details in the TEM micrographs. Interestingly, many of these undetermined cubic membrane morphologies are reported in pathological conditions in hominoidae.

3.2. Organelles with cubic membrane structure 3.2.1. Endoplasmic reticulum The ER was found to be the most prominent target of morphological alterations because of its highly convoluted and dynamic structure and crucial functions in membrane lipid synthesis and assembly, protein synthesis and secretion, ion homeostasis, and membrane quality control. These morphologies appear under numerous nicknames in the literature, such as ‘‘undulating membranes’’ (Schaff et al., 1976), ‘‘cotte de mailles’’ (Franke and Scheer, 1971), ‘‘membrane lattice’’ (Linder and Staehelin, 1980), ‘‘crystalloid membranes’’ (Yamamoto et al., 1996), ‘‘paracrystalline ER’’ (Wolf and Motzko, 1995), ‘‘tubuloreticular structures (TRS)’’ (Grimley and Schaff, 1976), and recently, as OSER (Snapp et al., 2003). Periodic symmetrical transitions of the ER are usually correlated with overexpression of certain ER-resident membrane proteins (Table 6.2) (see below). For example, overexpression of HMG-CoA reductase isozymes induces assembly of nuclear and cortical ER stacks with 2D symmetry, termed ‘‘karmellae’’ in yeast (Profant et al., 2000; Wright et al., 1988). Overexpression of this enzyme in UT-1 (Chin et al., 1982) or Chinese Hamster Ovary (CHO) cells (Jingami et al., 1987; Roitelman et al., 1992) induces formation of crystalloid ER, which houses most of the HMG-CoA reductase enzyme (Anderson et al., 1983; Orci et al., 1984). This correlation may imply a specific structure–function relationship of cubic membrane formation as a consequence of an altered protein or lipid inventory of the membrane. The cells of the phloem in plants are involved in the long-distance transport of nutrients and are known as sieve elements. Interestingly, the ER of differentiating sieve elements is a rare example in Nature in which

Table 6.2 Occurrence of crystalloid ER membranes in cell lines overexpressing certain ER-resident membrane proteins Description of cells/tissue

Overexpressed proteins

Cognomes

UT-1 cells (Compactin resistant CHO cells)

HMG-CoA reductase

Crystalloid ER

Hexagonal, cubic (G2)

CHO cells Yeast Yeast CV-1, COS-7

HMG-CoA reductase HMG-CoA reductase Cytochrome b(5) Cytochrome b(5)

Crystalloid ER Karmellae Karmellae Organized SER

COS-1 cells COS cells

msALDH InsP3 receptor

Crystalloid ER Cisternal stacks Tubular network

Hexagonal Multilayer lamellar Multilayer lamellar Multilayer lamellar (whorls), cubic (D2, G2) Cubic (G) Multilayer lamellar, whorls Retiform

Crystalloid ER

Hexagonal

Sandig et al. (1999)

Intracellular membrane Tubule

Hexagonal

Arechaga et al. (2000); Gales et al. (2002) Weiner et al. (1984)

CHO cells

Unassembled rubella virus E1 glycoprotein subunits HEK293 cells/ Cytochrome human P450 2B1 Escherichia coli Subunit (b) of F1F0 ATP synthase E. coli Fumarate reductase

Membrane organization

Hexagonal

References

Chin et al. (1982); Anderson et al. (1983); Pathak et al. (1986); Kochevar and Anderson (1987); Orci et al. (1984) Jingami et al. (1987) Wright et al. (1988) Verge`res et al. (1993) Snapp et al. (2003) Yamamoto et al. (1996) Takei et al. (1994) Hobman et al. (1992)

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two cubic membranes with the same structure but with different unit cell parameters, may coexist in the same cell (Behnke, 1965, 1968; Landh, 1996). These cells lack a nucleus and the cytoplasmic connection and exchange between vertically stacked cells is enabled through the perforated walls of the sieve elements (Behnke, 1965). Their function is to transfer the products of photosynthesis from the manufacturing site (leaves) to the storage cells (stem, roots, and seeds). The different types of cubic membranes may perhaps facilitate transport of various materials at different rates within the same cell. 3.2.2. Inner mitochondrial membranes Numerous researchers have reported mitochondria with inner membrane configurations that resemble cubic membrane morphologies (Brandt and Pappas, 1959; Kalt, 1974; Tahmisian et al., 1956). Possibly the bestcharacterized cubic membrane transition was observed in the mitochondrial inner membranes of the free-living giant amoeba, C. carolinense (Deng and Mieczkowski, 1998). In this organism, mitochondrial inner membranes undergo dramatic changes in 3D organization upon food depletion, providing an attractive, reversible model system to investigate induced membrane reorganization. Within one day of starvation, 70% of mitochondria undergo this morphological transition, which is observed in virtually all mitochondria after 7 days of starvation (Daniels and Breyer, 1968). This structural alteration of mitochondria in C. carolinense has been identified by a number of laboratories; however, in several reports the inner mitochondrial membranes take the appearance of tubular-like configurations that may appear in conjunction with well defined cubic membranes (Borysko and Roslansky, 1959; Brandt and Pappas, 1959; Daniels and Breyer, 1968; Sedar and Rudzinska, 1956). Indeed, by EM tomography, we have unambiguously demonstrated that inner mitochondrial membranes in C. carolinense cells adopt cubic morphology under starvation conditions (Deng et al., 1999). This induced transition is accompanied by alterations in cellular oxidative stress response, which led us to speculate that cubic membrane formation may be associated with oxidative stress (Deng et al., 2002) (see also discussion below). Intriguingly, formation of cubic membranes in amoeba Chaos is fully reversible to wild-type tubular morphology, upon refeeding (Deng and Mieczkowski, 1998). A similar cubic architecture of inner mitochondrial membranes was identified in a TEM ultrastructural study (Kalt, 1974) that describes mitochondrial pleomorphism in supporting (sustentacular) cells in testis of African clawed frog, Xenopus laevis. The mitochondrial membrane constellation in the mature stage of Sertoli sustentacular cells exhibits the D subtype of cubic membrane morphology. The mitochondria in the inner segment of the retinal cones of tree shrew species, Scandentia, the common tree shrew (Tupaia glis), and the northern

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tree shrew (Tupaia belangeri) (Foelix et al., 1987; Knabe and Kuhn, 1996; Knabe et al., 1997) are unique in size and ultrastructural arrangement of their inner membranes (Samorajski et al., 1966). These unusually large, patterned mitochondria exhibit one of the most complicated cubic membrane architectures known to date, with the highest possible symmetry (G subtype) of up to 12 layers of three-dimensionally folded membranes (see discussion below) (Fig. 6.7). 3.2.3. Plasma membrane Convoluted invaginations of the plasma membrane that are associated with the ‘‘smooth spongiome,’’ which is part of the contractile vacuole complex in Paramecium multimicronucleatum, have been proposed to be of the G subtype of cubic membrane organization (Allen, 2000; Landh, 1996; Patterson, 1980). Paramecium cells are able to maintain an almost constant intracellular osmolarity regardless of the environmental osmolarity. The complex regulation of the cell volume and osmotic gradient is primarily established by the smooth spongiome, which exhibits cubic membrane organization. Therefore, cubic membranes have been suggested to play roles in water segregation and intracellular volume and osmolarity control (Landh, 1996). ‘‘Honeycomb’’ structures of the T-tubular system in skeletal muscles have been observed in numerous diseased as well as in experimentally induced cases. Such structures were shown to be in continuity with the extra-cellular space and can thus be regarded as extensions of plasma membrane invaginations. The significance of honeycomb t-tubules is, however, unknown (Mastaglia and Walton, 1992), and surprisingly few studies deal with their 3D organization. Despite the elegant study of Ishikawa (1968), the 3D structure of these ‘‘honeycombs’’ was resolved only 30 years after their first discovery (Landh, 1996) and unambiguously demonstrated to be of cubic membrane morphology of the gyroid (G) subtype. 3.2.4. Photosynthesis-associated cubic membranes The structure of photosynthetic membranes and their assembly during development have been extensively studied (Gunning, 1965; Gunning and Jagoe, 1967; Gunning and Steer, 1975). In bacteria, several publications report lattice-like membrane morphologies as a variation of the more common multilamellar (thylakoid) configuration. In vegetative, photosynthetically active cyanobacteria, Anabaena sp., Lang and Rae (1967) reported a ‘‘prolamellar-like lattice’’, which was formed through continuous foldings of the photosynthetic thylakoid membranes. This cubic membrane bears a close resemblance to the prolamellar body (PLB) in higher plants, with the important difference that PLB are formed in the absence of light, whereas

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cubic membranes in Anabaena sp. were observed in fully illuminated cultures. The analysis of micrographs (Lang, 1965) of developing heterocysts (cells involved in nitrogen fixation) of Anabaena azollae clearly revealed several TEM projections of more or less developed cubic membranes (Landh, 1996). Although heterocysts lack the photosynthetic apparatus, these cubic membranes appear to arise by continuous folding of the thylakoid membranes. 3.2.4.1. Chloroplasts of green algae Zygnema In certain green algae species the chloroplast membrane(s) tend to form more complex morphologies than the simple ‘‘lamellar-like’’ structures. Chloroplasts of the green alga Zygnema transform to G-type cubic membrane (Fig. 6.6) during the log phase of cell culture (Deng and Landh, 1995). An analysis of previously reported electron micrographs by McLean and Pessoney (1970), which described ‘‘lamellar lattices’’ in Zygnema chloroplasts, indeed revealed a primitive (P) subtype of cubic membrane (Landh, 1996). This particular structure represents a continuous cubic membrane organization that is composed of several, mostly parallel, lipid bilayers (Deng, 1998). 3.2.4.2. Prolamellar body The fine structure of the PLB has been extensively studied by electron microscopy (Gunning, 1965; Gunning and Jagoe, 1967; Gunning and Steer, 1975; Israelachvili and Wolfe, 1980; Menke, 1962, 1963; Murakami et al., 1985; Osumi et al., 1984; Wehrmeyer, 1965a, b,c). Three basic space-lattice structures of the PLB have been proposed: (i) a primitive cubic lattice in which the fundamental unit is a hexapode (Granick, 1961; Gunning, 1965; Gunning and Jagoe, 1967), (ii) a diamond-based face-centered cubic lattice in which the fundamental unit is a tetrapode (Gunning and Steer, 1975; Murakami et al., 1985; Osumi et al., 1984; Wehrmeyer, 1965b), and (iii) a hexagonal Wurtzite-type of lattice (Ikeda, 1968; Wehrmeyer, 1965c; Weier and Brown, 1970). The double diamond-type cubic lattice is presumably the most common cubic structure of the PLB (Landh, 1996).

3.2.5. Inner nuclear membrane The formation of cubic membranes in the NE is not confined to the cytoplasmic side (i.e., the nuclear ER), but may also occur at intranuclear sites. This may appear surprising, however, formation of cubic membranes within nuclei could be realized by invaginations of a proliferating inner nuclear membrane. Intranuclear cubic or tubular membrane organizations usually develop in fast replicating neoplastic (tumor) cells (Babai et al., 1969; Karasaki, 1970) or non-neoplastic cells such as oocytes (Kessel and Beams, 1968; Miller, 1966). Their presence might play a role in mitotic activity or in facilitating nuclear-cytoplasmic communication. In human cells, intranuclear tubular (hexagonal or cubic) membrane organization was demonstrated in the endometrium during the secretory phase of the menstrual

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cycle (Bourgeois and Hubert, 1988; Ghadially, 1988). Progesterone and medroxyprogesterone can induce tubular structure formation in human endometrium (Kohorn et al., 1972), which suggests a role of an extrinsic hormonal factor in cell membrane organization.

4. Biogenesis of Cubic Membranes An extensive review of the literature reveals that virtually all membranes are capable of forming cubic structures. Their origin thus seems to be strongly coupled to the mechanisms of membrane biogenesis in general and, therefore, cubic membranes are to be considered as a defined configuration of cytomembranes. The formation and growth of cubic membranes may be a selective process to fulfill a specialized purpose under ever changing intracellular conditions. Thus, understanding the underlying rules(s) for the physiological selection between the different membrane morphologies is key. It is now well established that proteins may induce phase transition in lipid membranes resulting in new structures not observed in pure lipid– water systems (Bouligand, 1990). However, in principle, any amphiphilic molecule may be able to induce cubic membrane structures. Depending on the structure and nature of the proteins, their interactions with lipid bilayers can be manifested in very different ways. On the other hand, evidence from in vitro studies clearly differentiates membrane-forming lipids for their propensity to form nonlamellar structures, according to the molecular shape concept. The important structural role of membrane lipids in promoting cubic membrane formation in vivo is undisputed; however, only very recent evidence obtained from amoeba Chaos cells has correlated specific alterations of membrane lipids to cubic membrane formation in vivo (Deng et al., submitted). Although lipids extracted from these cells may assemble to cubic phases also in vitro (Fig. 6.8), marked differences in lattice size clearly indicate that additional factors—presumably proteins— exist in vivo that determine the overall 3D appearance of these structures. Induction of cubic membranes in Chaos cells upon starvation represents one rare example to experimentally address the molecular mechanisms leading to their formation in the biological context. From a topological point of view, cubic membranes appear to be formed from a structural ‘‘template’’ (the precursor of a cubic membrane), such as invaginations of a membrane. After initiation, further accumulation of membrane lipids may lead to intersection-free highly convoluted invaginations. During the folding process, the membrane must remain a continuous fluid structure that grows and interconnects without losing its polarity and integrity. Thus, the topology of the cubic membrane depends on the topology of its precursor structure. If one isolated invagination process

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A

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Figure 6.8 Lipid dispersion prepared from amoeba lipids (Deng, unpublished). TEM images of liposomes derived from lipids extracted from fed and 7d starved Chaos cells. (A) Multilamellar or whorl-like structures generated from fed cell lipids with numerous randomly distributed tubular structures, but without higher order phases. In contrast, (B) TEM data of lipid dispersion generated from lipids that were isolated from 7d starved amoeba cells show highly ordered domains.

triggers the membrane folding process, the topology will be that of a sphere, as is the case for vesicle formation during endocytosis or secretion. If more than one invagination takes place, it requires points of fusion to achieve a three-dimensionally interconnected membrane. Perhaps it is this symmetry that is the driving force for fusion; if fusion did not occur one would expect several independent cubic membrane systems to form, which would not necessarily bear any spatio-temporal correlation between each other or the periodicity of the template.

4.1. Role of membrane-resident proteins in cubic membrane formation Cubic membrane formation is frequently associated with the overexpression of certain ER-resident proteins and, to a lesser extent, with overexpression of some inner mitochondrial membrane proteins. Table 6.2 lists major membrane proteins that have been shown to induce cubic membrane formation upon experimental dys-regulation. 4.1.1. ER proteins HMG-CoA reductase is an ER-resident protein that is anchored to the membrane by seven membrane-spanning domains in its N-terminal part and has its catalytic domain extending to the cytoplasmic side. Elevated expression of HMG-CoA reductase is often associated with structural membrane alterations. The transmembrane region is indeed required to form crystalloid membrane structures of hexagonal (Fig. 6.5D) or cubic

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(Fig. 6.5C) morphologies, upon overexpression. Deletion of two of the seven membrane spanning regions or a truncated protein did not result in crystalloid ER formation, and the protein localized to disordered sheets rather than packed membrane tubules under these conditions. High expression levels of the soluble fragment of HMG-CoA reductase did not induce any crystalloid ER, again indicating that it is the transmembrane domain of HMG-CoA reductase that plays an important role in determining the structure of crystalloid ER ( Jingami et al., 1987; Yamamoto et al., 1996). Crystalloid ER is frequently observed in CHO cells upon overexpression of the HMG-CoA reductase gene, or in UT-1 cells, which are a mutant variant of CHO cells that overexpress this gene by 500 fold (Fig. 6.5; Chin et al., 1982). Notably, despite the presence of elevated levels of HMG-CoA reductase, which is the key enzyme of sterol biosynthesis, the membrane of the crystalloid ER appears to have very little cholesterol. Upon addition of cholesterol to UT-1 cells, which intercalates into the ER membrane, HMG-CoA reductase was subsequently degraded and the crystalloid ER disappeared ( Jingami et. al., 1987); sterol supplementation drastically reduced the rate of HMG-CoA reductase synthesis and also prevented the formation of new crystalloid ER. It was therefore speculated that the cubic membrane is an alteration in the feedback control of cholesterol synthesis, for the production of sterols and the biogenesis of smooth ER. Interestingly, administration of compactin, which is an HMG-CoA reductase inhibitor, also leads to HMG-CoA reductase overexpression, and induces the formation of stacked and aggregated structures, which were termed ‘‘karmellae’’ in yeast (Wright et al., 1988). The expression of msALDH in COS-1 cells also leads to alterations of the ER structure. Both HMG-CoA reductase and msALDH proteins possess large domains exposed on the cytoplasmic surface of the ER membrane, similar to the ER-resident protein, cytochrome P450. This led to the hypothesis that the formation of crystalloid membranes may require the expression of ER-resident proteins with large cytoplasmic domains (Sandig et al., 1999; Snapp et al., 2003; Yamamoto et al., 1996). Overexpression of specific ER-resident proteins such as cytochrome b (5) in COS-7 cells also triggers the formation of ‘‘whorls and crystalloid OSER structures’’ (Snapp et al., 2003). It was proposed that the biogenesis of OSER structures involves weak homotypic interactions between cytoplasmic domains of proteins and may underlie the formation of other stacked membrane structures within the cells as well. Time-lapse imaging of OSER biogenesis revealed that these structures formed rather quickly once a threshold level of OSER-inducing proteins was exceeded; OSERformation also involved gross remodeling of surrounding tubular ER.

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In this system, the attachment to the cytoplasmic domain of different ER-resident membrane proteins of green fluorescent protein (GFP) that is capable of low affinity, head-to-tail dimerization was sufficient to induce OSER formation upon overexpression in living cells. Homotypic low affinity interactions between cytoplasmic domains of proteins thus can differentiate tubular ER into stacked lamellae or crystalloid structures; such a mechanism may underlie the reorganization of other organelles into stacked structures as well (Snapp et al., 2003), and provides an intriguing model system to investigate the cellular and molecular requirements for cubic membrane formation. 4.1.2. Mitochondrial proteins Only a few mitochondrial proteins have been reported to induce and maintain tubular inner membrane morphology upon overexpression (Mannella, 2006), however, none of them was correlated with well-defined cubic membrane formation. Mitofilin, F1F0-ATPsynthase, and fission-fusion proteins may induce tubular or stacked lamellar and whorl-type membrane structures upon experimental overexpression. F1F0-ATPsynthase is an essential enzymatic complex of the mitochondrial inner membrane, which couples the proton electrochemical gradient generated by the respiratory chain to ATP synthesis. Various studies have shown that this complex is strongly implicated in the curvature of the inner mitochondrial membrane (reviewed by Voeltz and Prinz, 2007). Dimerization of the complex drives tubulation of the cristae (Dudkina et al., 2005; Strauss et al., 2008), while further oligomerization of these dimers is responsible for the formation and/or stabilization of inner membrane tubules. Mitofilin is a mitochondrial inner membrane protein, which assembles into a large multimeric protein complex. siRNA knockdown of mitofilin in HeLa cells yielded mitochondria with disorganized mitochondrial inner membranes: they failed to form tubular or vesicular cristae and appeared as intermittently fused, closely packed stacks of membrane sheets, resulting in a complex maze of membraneous networks (John et al., 2005). Mitofilin thus appears to be a key organizer of mitochondrial cristae morphology ( John et al., 2005). The role of mitochondrial proteins in cubic membrane formation in starved amoeba Chaos, which is a suitable model for analyzing reversible cubic membrane formation, is currently under investigation in the authors’ laboratories. 4.1.3. Morphogenic proteins Recently a class of membrane proteins known as morphogenic proteins was identified to shape the tubular ER in yeast and mammalian cells. These proteins are highly enriched in the tubular portions of the ER and virtually excluded from other regions. The study by Voeltz and coworkers (2006)

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illustrated the role of Rtn4a/NogoA, a member of the ubiquitous reticulon protein family that share a conserved C-terminal reticulon domain. Overexpression of Rtn4a/NogoA in mammalian cells, promoted the formation of ER tubules; membrane tubule formation in vitro, on the other hand, was prevented by anti-Rtn4a/NogoA antibodies. Similar results were observed in the yeast Saccharomyces cerevisiae, in which overexpression of Rtn1 (the yeast ortholog of Rtn4a/NogoA in mammals) also enhanced tubular ER formation. Yeast mutants lacking both Yop1 (an Rtn1 paralog) and Rtn1 showed a disrupted tubular ER, underscoring the important function of these proteins in shaping membrane structures. Reticulons contain long, hydrophobic domains that are inserted into the outer leaflet of the lipid bilayer. Since the hydrophobic domains are longer than required for spanning a bilayer membrane, it is believed that they promote a hairpin-like insertion into the lipid bilayer and give the overall appearance of a wedgelike protein. Thus, a local concentration of reticulons might induce and stabilize a high, positive membrane curvature. Although morphogenic proteins are believed to induce membrane curvature and shape spherical or tubular morphology, to date, none of them has been reported to induce highly organized membrane structures such as hexagonal or cubic membranes in vivo. One exception to the rule is the observation of a t-tubular system in skeletal muscles (Ishikawa, 1968), in which cubic membrane organization has been associated with caveolin-3 expression (Parton et al., 1997). Caveolin is synthesized in the ER but mostly resides in certain domains of the plasma membrane. Caveolins are known to play a role in inducing and maintaining membrane curvature. Individual caveolin molecules are cotranslationally integrated into the bilayer of the ER. Similar to reticulons, caveolins form a hairpin-loop within the bilayer, and tend to form hexa- or heptamers. These oligomers may leave from the ER and are transported via the Golgi apparatus to the plasma membrane, where, by a yet unknown mechanism, they induce localized sites of membrane curvature, known as caveolae (Bauer and Pelkmans, 2006; Voeltz and Prinz, 2007).

4.2. Role of lipids in cubic membrane formation Up to date, cubic membrane formation is mainly associated with overexpression of certain membrane-resident proteins. Although cubic phases are formed by ‘‘nonlamellar’’ lipids in vitro, only very few data directly correlate cubic membrane formation with cellular lipid profiles in vivo (Ryberg et al., 1983). In this study it was shown that the molar ratio of monogalactosyl diacylglycerol to digalactosyl diacylglycerol was higher in the PLB fraction that exhibits cubic membrane morphology, than in the prothylakoid fraction. Also, the content of glycolipids and protochlorophyllides was increased in the PLB fraction (Ryberg et al., 1983).

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Lipid analyses of fed and starved amoeba (C. carolinense) recently performed in the authors’ laboratory (Deng et al., submitted) may provide a first clue towards understanding the role of membrane lipids in determining cell membrane architecture. Detailed lipid analysis of amoeba Chaos exhibiting cubic membrane organization in their mitochondria, revealed an unusually high concentration of highly polyunsaturated fatty acids (C22:5; docosapentaenoic acid, DPA). Three predominant lipid species, namely plasmalogen PE (C16:0p/C22:5), plasmalogen PC (C16:0p/C22:5), and diacyl-PI (C22:5/C22:5), were identified in amoeba Chaos lipid extracts (Fig. 6.9), and their relative amounts increased up to 2.5-fold under starvation stress conditions (Deng et al., submitted). A rich body of data (for review see Nagan and Zoeller, 2001) suggests that plasmalogens—which are also present in mammalian cell membranes—may serve as mediators of membrane dynamics due to their high propensity to form inverted hexagonal structures. This property has also suggested a potential role for plasmalogens in facilitating membrane fusion processes (for review see Brites et al., 2004). Biophysical studies have also shown that the presence of plasmalogen PE lowers the lamellar to hexagonal-phase transition temperature (Lohner, 1996). Interestingly, CHO cells, which display massive membrane rearrangements upon HMG-CoA reductase overexpression (Fig. 6.5), are also rich N+

O

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Figure 6.9 Chemical structures of three major lipids found in membrane lipids extracted from amoeba C. carolinense: plasmologen PC (16:0p/22:5), plasmologen PE (16:0p/22:5), and diacyl PI (22:5/22:5).

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in plasmalogen lipids (up to 11% of their total phospholipids), especially plasmalogen PE (Nagan et al., 1998).

4.3. Electrostatic effects on cubic membrane organization The observation that weak molecular interactions, for example, by membrane proteins, may lead to cubic membrane transformation also suggests that electrostatic interactions play an important role in that process (Masum et al., 2005, Snapp et al., 2003). In a recent experimental study on the effects of divalent cations in the isolation buffers for mitochondria, we were able to demonstrate that the presence of EDTA up to 10 mM preserved mitochondrial cubic membranes in vitro (Fig. 6.10) (manuscript in preparation). Thus, pH and the nature of the electrolytes in the cellular milieu are likely to be important factors affecting cubic membrane morphology and integrity. In vitro studies using monoolein membranes containing negativelycharged dioleoylphosphatidic acid have shown that the electrostatic interactions caused by surface charge of the membranes (Li et al., 2001), charged short peptides such as poly-L-lysine (Masum et al., 2005), Ca2+ concentration (Awad et al., 2005), and low pH (Okamoto et al., 2008) play an important role in the phase transition between lamellar and cubic phases as well as on the stability of cubic phases. As a consequence of altered electrostatic interactions at the membrane interface, induced either by the A

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Figure 6.10 Electrostatic effects on cubic membrane organization (Deng, unpublished). Mitochondria of amoeba Chaos exhibiting cubic membrane arrangements were isolated in a buffer media containing (A) 50 mM, (B) 1 mM, or (C) 10 mM EDTA. Increasing the concentration of EDTA stabilized mitochondria with cubic morphology, suggesting a modulatory function of divalent cations in cubic membrane formation.

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increase in surface charge density of the membrane or by a decrease in salt concentration, the lipid membrane phase may change from the lamellar to the cubic phase. Such cubic phases may not only form to adopt the altered membrane proteins or charge distribution, but may also provide the means to rapidly adjust cellular physiology to the changing environmental conditions, such as fluctuations of (local) intracellular Ca2+ levels. The ‘‘breathing’’ of cubic membranes, that is, the changes in lattice size as demonstrated in vitro (de Campo et al., 2004), may represent an immediate biophysical response, which may allow for a rapid adaptation to the water content and ion concentration or charge distribution in a given membrane compartment also in vivo.

5. Cubic Membranes: Indicators of Cellular Stress and Disease? Notably, cubic membrane morphologies are often associated with deregulated protein synthesis, cellular stress, or more severe pathological conditions. This view, however, is perhaps somewhat biased since morphological abnormalities are typically investigated for diseased cells rather than for ‘‘wild type’’ conditions. Nevertheless, the correlation of specific membrane alterations in the course of acute or chronic cellular stresses needs closer consideration.

5.1. Virus-infected cells Virus-induced membrane transitions, generally referred to as cytomembraneous inclusions, are a hallmark in experimental and pathological samples (Almsherqi et al., 2005). However, due to the absence of a clear view of the 3D nature of these membrane structures, they appear under numerous nicknames in the literature, such as ‘‘tubulocrystalline inclusions’’ in HCV infected liver (Schaff et al., 1992), ‘‘convoluted membraneous mass’’ in viral St. Louis Encephalitis (Murphy et al., 1968), and ‘‘TRS’’ in SIV (Kaup et al., 2005) and SARS-virus infected Vero cells (Goldsmith et al., 2004). The unexplained mechanism behind the formation of convoluted cubic membranes upon viral infection starts to unfold based on the structural similarity to crystalloid ER membranes induced by overexpression of HMG-CoA reductase in UT-1, or the parental CHO cells (Fig. 6.5), as well as in A-431 (human epidermal carcinoma) cells. Some evidence suggests that virus infection and deregulation of HMG-CoA reductase are mechanistically linked, resulting in similar membrane phenotypes. For example, West Nile Virus infection is associated with local cholesterol alterations of the plasma membrane of the host cell that leads to a sequestration of membraneous sites of viral replication (Mackenzie et al., 2007). Depletion

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of cholesterol in the plasma membrane triggers HMG-CoA reductase upregulation and consequently leads to the formation of a crystalloid (cubic or hexagonal) ER membrane. The virus-induced convoluted membrane was reported to be essential for viral replication and survival: administration of cholesterol that reverses the HMG-CoA reducase-induced convoluted membrane formation in the infected cells significantly suppresses viral replication (Mackenzie et al., 2007). A recent study has also shown that virus-induced membrane complexes might provide partial protection against the host immune response (Hoenen et al., 2007) and hence offer a site for efficient viral replication (Mackenzie et al., 2007) or facilitate nucleo-cytoplasmic transport of genetic material (Almsherqi et al., 2008).

5.2. Neoplasia Cubic membranes (often labeled as TRS) have been reported to occur in the cytoplasm of breast carcinoma (Seman et al., 1971), subcutaneous myxoma (Stoebner et al., 1972), malignant lymphoma (Uzman et al., 1971), in the nucleus of human osteosarcoma cells (Murray et al., 1983), alveolar cell carcinoma (Ghadially et al., 1985), and gastric adenocarcinoma (Caruso, 1991). Since viral infections frequently lead to cubic membrane formation, it would not be surprising to observe cubic membrane structures also in cases of virus-related tumors, such as leukemias, lymphomas, and in virus-induced hepatocellular carcinoma (Uzman et al., 1971). TRS appear to be nonspecific to tumor differentiation or malignancy as they develop in highly replicative undifferentiated cells as well as in differentiated tumor cells. Therefore, due to the limited number of reported cases of cubic membrane structure in neoplastic cells, they currently are not yet diagnostically useful in the classification of tumors or in clinical prognosis. However, TRS could be used as a marker for viral-induced neoplasia, for example, hairy cell leukemia (Mantovani et al., 1986).

5.3. Muscular dystrophy Several muscular and neuromuscular diseases are associated with periodic membrane transformation of t-tubules of striated muscles, usually referred to as honeycomb structures. T-tubular morphological membrane transformation has been described in denervated muscles (Gori, 1972; Madarame et al., 1986; Miledi and Slater, 1969; Pellegrino and Franzini, 1963), in the muscles with induced experimental myopathy (Macdonald and Engel, 1970), bupivacaine-induced myonecrosis (Huxley and Taylor, 1958), and in tenotomized muscles of the rat (Andreev and Wassilev, 1994). They have also been observed in various human muscles as a consequence of muscular diseases (Borg et al., 1989; Cornog and Gonatas, 1967; Engel and Dale, 1968; Engel and Macdonald, 1970; Gori, 1972; Iorio Di et al., 1989;

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Miike et al., 1984). The fact that cubic membrane organization appears under pathological conditions of myopathy support the notion that they may be formed in response to stress conditions, such as a decrease in cell volume in the case of muscular dystrophy. Thus, it could be speculated that the potential function of cubic membrane structures is to maintain cellular integrity by regulating the volume of the cell (organelle) and/or to accommodate stress-induced membrane-resident proteins. Interestingly, induced formation of honeycomb t-tubules may also be part of a developmental program as these membrane structures are also observed in tadpole tail muscles that spontaneously degenerate during metamorphosis (Sasaki et al., 1985). Frequent transformation of t-tubules into G-type cubic membrane (honeycomb) organization in regenerating muscle fibers post pathological or experimentally induced muscular dystrophy may reflect the adaptive reorganization of the membrane system in the regenerating cell.

5.4. Autoimmune disease Cubic membranes (including those labeled as TRS) have been frequently reported to occur under conditions and in cell types with an immunological background, for instance in diseases in which autoantibody production is a main factor in the pathogenesis. These include diffuse connective tissue diseases such as systemic and discoid lupus erythematosus (Grimley and Schaff, 1976) and Sjo¨gren syndrome (Nakamura, 1974) or organ limited diseases, such as thyroiditis, gastritis, myasthenia gravis, and celiac disease (Helder and Feltkamp-Vroom, 1974). TRS are also reported in autoimmune diseases characterized by deposition of immune complexes or factors, which are discrete in comparison to the diffuse connective tissue diseases like rheumatoid arthritis, nephritis, and amyloidosis (Helder and FeltkampVroom, 1974). Indeed, cubic membrane organization is frequently reported in immune deficiency diseases such as HIV (Kaup et al., 2005). Such ultrastructural membrane alterations are often reported in endothelial cells and in lymphocytes; therefore, TRS should be regarded as potential markers of autoimmune diseases that are of a systemic nature.

6. Cubic Membranes: Specific Functions or Innocent Bystanders? The frequent appearance of cubic membranes in several specialized cell types, in response to environmental conditions, stress or infection, poses the intriguing question as to their specific functions. It is possible that cubic membranes are but an inevitable self-assembled product of the complex molecular mixture of lipids and proteins, the result of ‘‘simple’’ molecular packing considerations and inter-molecular interactions. Even though this is

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appealing to the long and unresolved debate about ‘‘nonlamellar’’ lipids in conjunction with cell membranes, an increasing body of evidence suggests that these structural organizations might have to fulfill a purpose and their formation cannot be rationalized solely by spontaneous molecular packing. It is important to stress that the proposed functions of cubic membranes in the following discussion are hypothetical, although recent experimental data provide support to some of these hypotheses.

6.1. Cell space organization and subvolume regulation Continuous hyperbolic layers, as represented by cubic membranes, partition space into discrete and unconnected subvolumes. By definition, a multimembraneous structure consisting of (n) membranes, partitions space into (n þ 1) physically distinct, intertwined, but separate subspaces. The identity of each space itself is not a consequence of the existence of a cubic membrane; rather it is an effect of a topologically invariant membrane morphogenesis. Although an intracellular space maintains its identity as long as membrane continuity is preserved, it is only by means of a hyperbolic membrane structure that such spatial relations can be rigorously defined. The identification of cubic membranes in multiple cell types and tissues clearly demonstrates that continuous intracellular membranes are much more frequently adopted than generally acknowledged.

6.2. Inter- and intracellular trafficking Intracellular and inter-organelle trafficking of macromolecules and communication are active areas of research, yet no unified model exists. To what extent intracellular trafficking depends on or can be optimized through specific types of membrane organization, for example, as continuous 3D space partitioners or networks, is unknown. In the three dimensions of a cell, the partitioning of the intracellular space into subvolumes—leading to an interpenetrated pipeline system—could in principle be orchestrated by membrane organization, which ultimately controls spatio-temporal activities, including intracellular trafficking and transportation, and fast response to physiological alterations such as changing temperature, osmolarity or pH. Recognition of the existence of topologically distinct spaces through the formation of cubic membranes implies that cell space is restricted and predetermined to maintain distinct but interconnected subcellular domains. Cubic membranes may induce enrichment—or exclusion—of certain lipids and proteins, as a prerequisite for vesicle-mediated subcellular trafficking. They may also function as partitioners within a membrane, to create subdomains of specific molecular composition and function. It has been repeatedly noted that the stage of cell division and differentiation has a strong influence on the formation of cubic membranes. Cells that undergo rapid cell division and

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differentiation, such as the differentiating sieve elements, cells during spermatogenesis, or tumor cells are well over-represented in the collection of cubic-membrane forming cells (Table 6.1). Obviously, during cell division and differentiation events, there is a greater need for regulated spatio-temporal transport, as well as cellular communication, which is particularly well illustrated during spermatogenesis (Table 6.1). In the case of differentiating sieve elements, cubic membranes might have a particular role in the assimilated transport process. Finally, it is of interest to note that aggregates of ‘‘synaptic vesicles’’ often resemble cubic membranes. This can be taken as an indication of a possible on-off mechanism of membrane continuity, which might account for a regulative capacity for the controlled release of transmitter substances.

6.3. Specific structure-function relationships Several more general functionalities are under consideration for cubic membranes. In particular, a cubic bilayer arrangement could serve as a regulator of chemical or physical potentials across the membrane surface. This is indicated in several cases in which cubic membranes have been found to be sensitive to the growth medium. An osmo-regulative capacity of a plasma membrane-associated cubic membrane can be envisioned, which might serve to allow rapid adjustment to osmotic changes. Such a function is, for instance, consistent with the presence of cubic membranes in a variety of epithelial cells (Table 6.1). Cubic membranes may also be involved in curvature-controlled activation and regulation of membrane enzyme activities. For instance, diacylglycerols, which may function as second messengers to activate protein kinase C (PKC), display a strong propensity to form negatively curved membrane structures. The activity of PKC, however, is not specifically dependent on lipids that induce reverse hexagonal phases (Senisterra and Epand, 1993), and thus, reversed cubic phases. In view of the ubiquitous occurrence of cubic membranes it would be of interest to investigate the influence of PKC on cubic phases, and vice versa, to eventually correlate the geometry of the environment to protein activity. Certain cubic membrane structures are apparently strongly linked to specific functions, such as the cubic structure found in the PLB of photosynthetic cells in higher plants. A physical role has been proposed for the selection of D-surface geometry (Guo et al., 1995), based on theoretical considerations on how the geometry of crystals controls the emission or absorption of certain wavelengths of light through the existence of photonic band-gaps (Babin et al., 2002). Although the D-surface is isotropic, its geometry is such that it may potentially trap photons. The absorbed energy could then be used by certain molecules that are positioned along a particular lattice direction, and, for instance, trigger conformational changes to revert PLB into an active thylakoid membrane geometry. A similar structural role can be considered for the selection of different subtypes of cubic

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membrane architecture (photosome) in bioluminescent scaleworms (Bassot, 1964, 1966). Interestingly, the photosome membrane organization has a D subtype cubic membrane in the resting (unstimulated) state, however, upon electrical or optical stimulation the membrane configuration changes and acquires P subtype. Do different cubic membrane subtypes manipulate different wavelengths? Does the function of the cubic membrane change with the specific alteration of its geometrical configuration? The answers to these questions clearly deserve further investigations. In addition, several other photoactive cell types exist that contain cubic membranes. Mitochondria in the retinal cone of the tree shrew (Tupaia glis) (Foelix et al., 1987) may adopt an isotropic membrane structure that allows efficient capture of the incoming light; in an alternative model, this multilayer G-surface cubic arrangement of the mitochondrial membrane (Fig. 6.7) may preferentially reflect UV light before reaching the outer segment of the retina, thus representing a specialized optical filter system.

7. Applications of Cubic Membranes Although the specific functions of cubic membranes in biological systems are still to be uncovered, recent studies on the physical properties of such highly ordered membrane structures have suggested a range of potential applications, beyond conventional liposome technologies. Dispersed liquid crystalline phases such as bicontinuous cubic dispersions (known as CubosomesÒ ) have been the focus of recent studies as potential drug delivery agents (Barauskas et al., 2005). Lipids forming cubic phases in vitro are also well-established matrices, for example, for membrane protein crystallization (Seddon et al., 2004), but are not considered here in the context of cubic membranes.

7.1. DNA transfection Cubic membrane function is potentially linked to intracellular trafficking processes for macromolecules. Indeed, isolated mitochondria with cubic membrane organization from starved amoeba cells (see above) have recently been shown to incorporate and retain short oligonucleotides (ODN) with high efficiency (Fig. 6.11) (Almsherqi et al., 2008). Although the molecular basis for the high affinity of ODN to cubic membranes has not been uncovered yet, it is reasonable to assume that it is strongly promoted by electrostatic interactions with the highly curved and extended membrane surface. Furthermore, delivery of ODN-cubic membrane complexes to mammalian cells in culture appears to be very effective and might, therefore, provide an attractive alternative to currently employed cationic

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A

B

0.5 µm

2 µm

D

C

2 µm

1 µm

Figure 6.11 Cubic membrane organization and DNA uptake (Almsherqi et al., 2008). (A) Low and (B) high magnification TEM images of mitochondria containing cubic membrane structure isolated from 10 d starved Chaos cells before (A and B) and after (C and D) incubation with ODNs. Multiple electron-dense intra-mitochondrial inclusions (D) may represent cubic membrane-mediated ODN interactions. The multiple pores (B) at the surface of mitochondria with cubic membrane organization may play an important role in facilitating passive uptake of ODNs.

lipoplexes and promises high efficacy and reduced cytotoxicity. Use of cubic membranes of biological origin as a carrier for DNA transfection or delivery of other nucleic acids, such as small interfering RNA and/or short hairpin RNA is currently under active investigation (Almsherqi et al., 2008).

7.2. Do cubic membranes have optical properties? The photonic properties of bi- or multicontinuous cubic phases based on triply periodic level surfaces (G, D, and P) have been extensively studied in materials science (Babin et al., 2002; Maldovan et al., 2002; Urbas et al., 2002). In general, the lattice size of the photonic crystal has to be of the same magnitude as the wavelength range it controls. Photonic crystals based on triply periodic surfaces with the largest photonic band gaps known are of type D and G cubic morphology (Babin et al., 2002). Similar G-surfaces describe

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the cubic membrane organization of the inner mitochondrial membranes (Fig. 6.7) in the tree shrew’s (Tupaia belangeri) retina (Foelix et al., 1987) and photosynthesis-related membranes in prolamellar bodies of higher plants and chloroplasts of the green algae Zygnema (Fig. 6.6) (Deng, 1998; Deng and Landh, 1995). Based on these similar cubic geometries and relevant lattice sizes, it is tempting to speculate that perhaps Tupaia mitochondrial membranes and cubic membranes in Zygnema chloroplasts and PLB of higher plants may function as photonic crystals. The 12-layered cubic membrane organization of the mitochondria in retinal cones of tree shrew Tupaia and multilayer grana of photosynthetic membranes may thus enable these membranes to amplify, refract, or absorb certain wavelengths of light. Whether such an elaborate membrane arrangement is a protective response or whether it provides a specialized reaction center for embedded proteins awaits detailed analysis.

8. Concluding Remarks The frequent appearance of nonlamellar membrane arrangements such as cubic membranes in cells under stressed or pathological conditions points to an intrinsic cellular response mechanisms. The extensive generation of membrane surfaces to facilitate exchange reactions between two (or more) sub-volumes, or the sequestration of specific lipids or proteins into membrane domains of nonlamellar morphology may provide a physiological advantage to cope with various stress phenomena. Current analyses of nonlamellar membrane arrangements are largely restricted to the descriptive level; the identification of inducible membrane systems, such as in virus-infected cells or the reversible transition of mitochondrial inner membranes of amoeba Chaos to cubic morphology upon starvation, however, opens new avenues for understanding the molecular mechanisms and cellular requirements underlying these membrane arrangements. Lipidomics and proteomics techniques on the one hand, and cryo-electron tomography on the other, hold great promise to uncover the mysteries of cubic membranes in living cells, in healthy and diseased states.

ACKNOWLEDGMENTS The authors apologize to the colleagues whose work has not been cited due to space limitations. We thank Felix Margadant for critical reading of the manuscript, Mark Mieczkowski for the ‘‘Cubic Membrane Simulation Projection’’ program (QMSP), and Aik Kia Khaw for his art work presented in Fig. 6.4. The technical support by Chwee Wah Low, Li Ling Olivia Tan, and Mei Yin Shoon is gratefully acknowledged. Research in the authors’ laboratories is supported by research grants NMRC (R-185-000-058-213) and BMRC from Singapore (R-185-000-065-305) to Y. D. and SFB Lipotox (project F3005) of the Austrian Science Fund to S.D.K.

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Index

A Abiotic stress response, 221 Actin microfilament stabilization expression-related phenotype changes, 31 hormone release control, 30 over expression, cDNA transfection, 30–31 AEE. See Apical early endosome Anabaena sp, 310–311 Apical early endosome (AEE), 132 Autoimmune disease, 321 Autophagy, 219 B Basolateral early endosome (BEE), 132 BDNF. See Brain-derived neurotrophic factor Biogenesis, cubic membranes cytomembranes, 312 electrostatic effects, 318–319 lipids, 316–318 membrane-resident proteins electrostatic effects, 318–319 ER proteins, 313–315 lipids, 316–318 mitochondrial proteins, 315 morphogenic proteins, 315–316 molecular shape concept, 312 Brain-derived neurotrophic factor (BDNF), 255 C Caldesmon (CaD) CALD1 gene expression myogenesis, 4–5 podosomes formation, 4 50 -splice site selection, 5 transcriptional regulation, 5–7 calponin interaction actin-myosin interaction, 15, 23 actomyosin ATPase inhibition, 23 h2-calponin, 23–24 myosin and actin-linked regulation, 15 property comparisons, 19–22 domain mapping Ca2þ–calmodulin binding sites, 13 CaD39 and CaD40, 8, 11 tropomyosin binding site, 11

gene organization CaD isoforms, 3 human CADL1 gene, 3–4 HeLa l-CaD expression, 43–44 intracellular localization anti-tropomyosin antibody injection, 29 platelet activation, 29–30 podosome formation, 28–29 stress fibers, 28 nonmuscle cells actin microfilament stabilization, 30–31 Ca2þ–calmodulin, 32–35 phosphorylation, 35–41 partial bladder outlet obstruction, 43 phosphorylated residues oncogenic v-erbB, 14–15 phosphorylated sites and effects, 16–17 podosome formation, cortactin CD44 receptor activation, 41–42 ERK phosphorylation, 42–43 N-WASP-mediated Arp2/3 activation, 42 smooth muscle contraction, h-CaD ERK and p38MAPK, 25–26 phosphorylation, 25–27 therapeutic strategy, 45–47 Cald1 gene HeLa type l-CaD, 43 organization, 3–4 podosomes formation, 4 regulation myogenesis, 4–5 50 -splice site selection, 4–5 transcriptional regulation, 5–7 Calponin interaction, Caldesmon actin-myosin interaction, 15, 23 actomyosin ATPase inhibition, 23 h2-calponin, 23–24 myosin and actin-linked regulation, 15 property comparisons, 19–22 Cancer stem cells (CSCs), 156 CDK. See Cystic diseases of kidney Cell architecture cell–cell junctions junctional complexes, 137–138 polarity complexes, 138–141 differentiated plasma membrane domains apical and basolateral membrane, 131 CRE, 131 membrane fusion machinery, 135–137

343

344 Cell architecture (cont.) sorting signals and machineries, 133–135 trafficking pathways, 131–132 trans-Golgi network (TGN), 131 Cell–cell junctions, epithelial tube formation junctional complexes apical juctions (AJs ), 137–138 tight junctions (TJs), 137 polarity complex apical–basolateral polarization, 140 LKB1, 138 mammalian cell, 138–139 signaling pathway, 139 CFTR. See Cystic fibrosis transmembrane conductance regulator Chaos carolinense, 302, 305, 309, 317 Choroideremia, 160, 253 Common recycling endosomes (CRE), 131 Cortactin, podosome formation CD44 receptor activation, 41–42 ERK phosphorylation, 42–43 N-WASP-mediated Arp2/3 activation, 42 CRE. See Common recycling endosomes CSCs. See Cancer stem cells Cubic membranes applications DNA transfection, 324–325 optical properties, 325–326 autoimmune disease, 321 biogenesis cytomembranes, 312 electrostatic effects, 318–319 lipids, 316–318 membrane-resident proteins, 313–316 molecular shape concept, 312 cell space organization and subvolume regulation, 322 inter-and intracellular trafficking, 322–323 membrane morphology, transmission electron microscopy cryo-ET, 305 direct template correlative (DTC) matching method, 305–306 electron tomography (ET), 305 membrane polymorphisms Chaos carolinense, 302 ER proteins, 302–303 membrane symmetries distinct morphological patterns, 279 fixation method, 281 patterned membrane organization, 279 tubulo-reticular structure (TRS), 280–301 muscular dystrophy, 320–321 neoplasia, 320 organelles endoplasmic reticulum (ER), 307–309 inner mitochondrial membranes, 309–310 plasma membrane, 310

Index

photosynthesis-associated membranes Anabaena sp, 310–311 inner nuclear membrane, 311–312 prolamellar body (PLB), 311 prolamellar-like lattice, 310 Zygnema, chloroplasts, 311 protozoa–mammals, 306–307 specific structure-function relationships, 323–324 virus-infected cells, 319–320 vs. cubic phases equilibrium process, 303 Zygnema sp., 303–304 Cystic diseases of kidney (CDK), 164–165 Cystic fibrosis (CF), 158 Cystic fibrosis transmembrane conductance regulator (CFTR), 158–159, 254 Cytokinesis RAB, 214–215 SNARE, 215 Cytotoxic T lymphocytes (CTL), 254 D De novo formation, epithelial tube central lumen antiadhesive/repulsive factors, 148 mechanism, 146–147 Slit–Robo signaling, 148 vacuolar apical compartment (VAC), 146 phosphoinositides, membrane identity PtdIns(3,4,5)p3, 151–152 PtdIns(4,5)p2, 149–151 polarity axis, orientaion cadherins and integrins, 144–146 3D-MDCK system, 145 extracellular matrix interaction, 144 GTPases, 146 Differentiated plasma membrane, polarized cell apical and basolateral membrane, 131 membrane fusion machinery annexin A2, 136 basolateral vesicle docking, 135–136 SNARE complex, 135, 137 sorting signals and machineries apical sorting, 133 basolateral sorting, 134–135 FAPP2, binding protein, 134 GPI anchor and lipid-raft hypothesis, 133 MARVEL domain, 133–134 transcytosis, 135 trafficking pathways, 131–132 trans-Golgi network (TGN), 131 DNA transfection, 324–325 E E-cadherin, 138, 153 EMT. See Epithelial-to-mesenchymal transition

345

Index

Endoplasmic reticulum (ER) cubic membrane polymorphisms, 302–303 membrane-resident proteins crystalloid membrane structures, 313–314 green fluorescent protein (GFP), 315 HMG-CoA reductase, 313–314 karmellae, 314 organelles, 307–309 Epithelial polarity cancer cells annexins, 156 E-cadherin, 153 EMT, 152–153 HPV strains, 153 neoplastic tissues, 152 PAR–aPKC complex, 153–154 phosphatase and tensin homolog, 154–155 stem cell theory, 156 three dimensional cultures, 156–157 VHL, 155 CDKs, 164–165 cytoskeletal and phosphoinositide disorder cystic diseases, kidney (CDKs), 164–165 GTPases and Rho regulators, 161–162 microvillus inclusion disease (MID), 162–163 phosphoinositide metabolism, 163–164 trafficking disorders adaptor protien disruption, 159–160 protein sorting signal, defect, 157–159 Rab GTPases, defect, 160–161 Epithelial-to-mesenchymal transition (EMT), 130, 152, 154–155 Epithelial tube formation cell architecture cell–cell junctions and polarity complexes, 137–141 differentiated plasma membrane, 131–137 de novo formation central lumen formation, 146–148 phosphoinositides (PtdIns) role, 149–152 polarity axis, 144–146 morphogenesis diverse patterns, 143–144 tubulogenesis, 141–143 polarity and disease CDKs, 164–165 cell polarity and cancer, 152–157 cytoskeletal and phosphoinositide disorder, 161–164 trafficking disorders, 157–161 structural and cellular diversity, 130 ER-Golgi trafficking ER, SNAREs, 206–207 Golgi apparatus, SNAREs, 207 RAB2/RABB and RAB6/RABH, 206 RAB1/RABD, 205–206 Exocyst protein complex, 135–136

G GABA-GnRH connection, HPG axis adulthood agonists/antagonists effects, 104 electrophysiological studies, 106–107 GAD67 mRNA expression, 100 GnRH gene expression, 102–103 GnRH/LH surge, 99–102 pulsatile LH release, 97–99 steroid regulation, 103–106 early development bicuculline application, 94 excitatory to inhibitory, 92–93 GABAAR protein expression, 93 3 H-muscimol binding, 94 puberty AOAA treatment, 97 depolarized GnRH neurons, 96 heterogenous population, GABAARs, 95 pubertal transition, 96–97 reproductive senescence, 107–108 Gamma-aminobutyric acid (GABA) receptors expression, 91–92 GABA–GnRH connection, 92–94 ionotropic receptor classes, 91 structural properties, 92 subtypes and GAD isoforms, 90 G-domain, 236 GEF. See Guanine-nucleotide exchange factor Glutamate–GABA interactions AOAA inhibitor, 111 developmental transition, 110 GABAAR and GABABR, 110–112 neuron structural model, 109–110 POA (preoptic area) examination, 111–112 P15 to P30 transition, 110–111 vesicular GABA transporter (VGAT), 91, 112 Glutamate–GnRH connection adulthood NMDAR and non-NMDAR effects, 84 reproductive physiology regulation, 86 steroid modulation, GnRH neurons, 84–85 early development, 78–79 puberty, 81–83 reproductive senescence age–related changes, receptor expression, 86–87 aging affects glutamate levels, 90 anteroventral peroventricular nucleus, 88 coexpression changes, 87–89 functional changes, 89 Glycosylphosphatidylinositol (GPI), 133 gp135. See Podocalyxin Gravitropism, 216–217 Griscelli syndrome (GS), 253 GT1–7 cell line, 77–78 GTPase inhibiting proteins (GIP), 242

346

Index

GTPase, Rab proteins C-terminal region, 246, 250 effectors dynein, 260–261 endogenous Rab6A and giantin, 262 mitotic kinesin-like protein2 (MKlp2), 259–260 Rab6A and Mint3, colocalization study, 261 western blotting, 263 guanosine diphosphate dissociation inhibitor, 244, 246 Huntington disease (HD), 255 malignancy brain-derived neurotrophic factor (BDNF), 255 cardiac hypertrophy, 254 inclusion membrane proteins (Incs), 256 Warburg micro syndrome and Martsolf syndrome, 253 nomenclature, subcellular localization and functions, 237–241 OCRL1 protein, 254 Rab escort protein (REP), 244–246 role and function, 237 structure and phylogeny carboxyterminal domains, 247 core fold, 246 isoforms, 252 Rab complementarity-determining region (RabCDR), 247 Rab/Ypt family, 252 Guanine-nucleotide exchange factor (GEF), 244–246 Guanosine diphosphate (GDP) dissociation inhibitor (GDI), 242, 246 H h-CaD, smooth muscle contraction ERK and p38MAPK, 25–26 phosphorylation, 25–27 HeLa l-CaD expression, 43–44 Hemaglutinin, 133 Hermansky–Pudlak syndrome (HPS), 159, 253 Higher order plant functions abiotic stress response, 221 autophagy, 219 cell differentiation, morphogenesis, and flowering phenomena, 221–222 gravitropism, 216–217 plant–microbe interaction, 220–221 tip growth, 217–219 HMG-CoA reductase, 313 HPS. See Hermansky–Pudlak syndrome Human Papillomavirus (HPV), 153 Huntington disease (HD), 255 Hypothalamic neural systems GABA-GnRH connection

adulthood, 97–107 early development, 92–94 puberty, 95–97 reproductive senescence, 107–108 GABA receptors expression, 91–92 GABA–GnRH connection, 92–94 ionotropic receptor classes, 91 structural properties, 92 subtypes and GAD isoforms, 90 GAD67 mRNA expression, 99–100, 105, 108 glutamate/GABA interactions developmental transition, 110 GABAAR and GABABR, 110–112 neuron structural model, 109–110 OAA inhibitor, 111 POA examination, 111–112 P15 to P30 transition, 110–111 vesicular GABA transporter (VGAT), 91, 112 glutamate-GnRH connection adulthood, 83–86 early development, 78–79 puberty, 79–83 reproductive senescence, 86–90 glutamate receptors, HPG axis AMPA-type receptors, 77 glutamatergic subtypes, 76 GnRH neurons, 75–76 GT1–7 cell line, binding studies, 78 immortalized GnRH cell line, GT1–7, 77 immunohistochemical analysis, 75 ionotropic receptor classes, 76 localization, 75 NMDAR, 76–77, 80 structural properties, 78 GnRH perikarya, 75, 77 reproductive life cycle, females critical biological rhythm, 72–73 estrous cycles, puberty, 72 GnRH–glutamate–GABA connection, 74–75 senescence, 73–74 sexual maturation, 71–72 Hypothalamic-pituitary-gonadal (HPG) axis. See also Hypothalamic neural systems GABA receptors expression, 91–92 GABA–GnRH connection, 92–94 ionotropic receptor classes, 91 structural properties, 92 subtypes and GAD isoforms, 90 glutamate–GnRH connection adulthood, 84–86 early development, 78–79 puberty, 81–83 reproductive senescence, 86–90 glutamate receptors

347

Index

AMPA-type receptors, 77 glutamatergic subtypes, 76 GnRH neurons, 75–76 GT1–7 cell line, binding studies, 78 immortalized GnRH cell line, GT1–7, 77 immunohistochemical analysis, 75 ionotropic receptor classes, 76 localization, 75 NMDAR, 76–77, 80 structural properties, 78 I Inner mitochondrial membranes, 309–310 Intracellular trafficking pathways A. thaliana, 203–204 cytokinesis RAB, 214–215 SNARE, 215 ER-Golgi trafficking ER, SNAREs, 206–207 Golgi apparatus, SNAREs, 207 RAB2/RABB and RAB6/RABH, 206 RAB1/RABD, 205–206 secretion plasma membrane, SNARE, 209–210 RAB11/RABA, 208–209 RAB8/RABE, 207–208 trans-Golgi-network (TGN) AtSYP41/ TLG2a and AtSYP42/TLG2b, 210–211 vacuolar and PVC /endosomal trafficking RAB18/RABC, 213 RAB5/RABF, 211–213 RAB7/RABG, 211 SNARE, 213–214 vesicular transport pathway, 203 Ionotropic glutamate receptors, 75–78 L Late endosomes (LE), 159 M Madin–Darby Canine Kidney (MDCK) cell, 136 Malignancy, Rab GTPases brain-derived neurotrophic factor (BDNF), 255 cardiac hypertrophy, 254 inclusion membrane proteins (Incs), 256 Warburg micro syndrome and Martsolf syndrome, 253 Martsolf syndrome, 253 Medial preoptic nucleus (MPN), 75 Membrane fusion machinery annexin A2, 136 basolateral vesicle docking, 135–136 SNARE complex, 135, 137

Membrane polymorphisms Chaos carolinense, 302 ER proteins, 302–303 Membrane-resident proteins electrostatic effects, 318–319 ER proteins crystalloid membrane structures, 313–314 green fluorescent protein (GFP), 315 HMG-CoA reductase, 313–314 karmellae, 314 lipids chemical structures, 317 molar ratio and glycolipid content, 316 plasmalogens, 317 mitochondrial proteins, 315 morphogenic proteins, 315–316 Membrane symmetry distinct morphological patterns, 279 fixation method, 281 patterned membrane organization, 279 tubulo-reticular structure (TRS), 280–301 Membrane trafficking, RAB and SNARE proteins A. thaliana, 199–200 downstream reactions, RAB effectors, 190–195 genome sequence, 199 GTPase cycle, 188–189 hermaphrodites, 198 N-ethyl-maleimide-sensitive factor attachment protein (SNAP), 187 plants, life style, 196–198 tethering and fusion machinery, SNARE molecules, 195–196 Microvillus inclusion disease (MID), 161–163 Morphogenesis diverse patterns, tube formation polarized epithelia, 143 unpolarized epithelia, 143–144 tubulogenesis in vitro system, 141–142 in vivo system, 142–143 MPN. See Medial preoptic nucleus Muscular dystrophy, 320–321 N Neoplasia, 320 NMDA and non-NMDA receptors, 75–85, 89, 110–112. See also Hypothalamic neural systems Non muscle cell, Caldesmon actin microfilament stabilization expression-related phenotype changes, 31 hormone release control, 30 over expression, cDNA transfection, 30–31 2þ Ca –calmodulin A7r5 cell line, 34

348

Index

Non muscle cell, Caldesmon (cont.) Arp2/3 complex, 34–35 CaD39-AB, 32–34 microinjection studies, 32 intracellular localization anti-tropomyosin antibody injection, 29 platelet activation, 29–30 podosome formation, 28–29 stress fibers, 28 l-CaD, phosphorylation Cdc2 kinase, 35–38 ERK, 38–39 PAK, 40–41 O Organelles endoplasmic reticulum (ER), 307–309 inner mitochondrial membranes, 309–310 plasma membrane, 310 P Paramecium multimicronucleatum, 310 Partial bladder outlet obstruction (PBOO), 43 Peutz–Jeghers syndrome (PJS), 138 Phosphatase and tensin homolog (PTEN), 149, 151 Phosphatidylinositol-4,5-bisphosphate (PtdIns (4,5)p2), 149–151 Phosphatidylinositol-3,4,5-trisphosphate (PtdIns (3,4,5)p3), 151–152 Phosphomannomutases (PMMs), 263–265 Photosynthesis-associated membranes Anabaena sp, 310–311 inner nuclear membrane, 311–312 prolamellar body (PLB), 311 prolamellar-like lattice, 310 Zygnema, chloroplasts, 311 Plant–microbe interaction plant immune system, 220 symbiosis and nodule formation, 220–221 Plasma membrane, 310 Podocalyxin, 148 Podosome formation, cortactin CD44 receptor activation, 41–42 ERK phosphorylation, 42–43 N-WASP-mediated Arp2/3 activation, 42 PTEN. See Phosphatase and tensin homolog Puberty, HPG axis GABA-GnRH connection AOAA treatment, 97 depolarized GnRH neurons, 96 functional changes, 89 heterogenous population, GABAARs, 95 pubertal transition, 96–97 glutamate-GnRH connection, 79–83

R RAB and SNARE proteins higher order plant functions abiotic stress response, 221 autophagy, 219 cell differentiation, morphogenesis, and flowering phenomena, 221–222 gravitropism, 216–217 plant–microbe interaction, 220–221 tip growth, 217–219 membrane trafficking A. thaliana, 199–200 downstream reactions, RAB effectors, 190–195 genome sequence, 199 GTPase cycle, 188–189 hermaphrodites, 198 N-ethyl-maleimide-sensitive factor attachment protein (SNAP), 187 plants, life style, 196–198 tethering and fusion machinery, SNARE molecules, 195–196 molecular/cellular research biochemistry, cytology, and forward genetics, 201–203 cytokinesis, 214–215 ER–Golgi pathways, 205–207 plants, identification, 200–201 plant vesicular trafficking, post genome era, 203 secretion, 207–211 vacuolar and PVC /endosomal trafficking, 211–214 organelles, 184 unique regulatory system, endocytic/vacuolar transport, 222–223 Rab complementarity-determining region (RabCDR), 247 Rab escort protein (REP), 244–245 RAB proteins molecular/cellular research A. thaliana, 200–201 biochemistry, cytology, and forward genetics, 201–202 cross-hybridization, 200 cytokinesis, 214 ER–Golgi, 205–206 secretion, 207–209 vacuolar and PVC /endosomal trafficking, 211–213 Rab proteins active and inactive mutants carboxyterminal domains, 247 RabF motifs, 247–250 GTPase brain-derived neurotrophic factor (BDNF), 255 carboxyterminal domains, 247

349

Index

cardiac hypertrophy, 254 core fold, 246 C-terminal region, 246, 250 dynein, 260–261 endogenous Rab6A and giantin, 262 guanosine diphosphate dissociation inhibitor, 244, 246 Huntington disease (HD), 255 inclusion membrane proteins (Incs), 256 isoforms, 252 mitotic kinesin-like protein2 (MKlp2), 258–260 nomenclature, subcellular localization and functions, 237–241 OCRL1 protein, 254 Rab6A and Mint3, colocalization study, 261 Rab complementarity-determining region (RabCDR), 247 Rab escort protein (REP), 244–246 Rab/Ypt family, 252 roles and functions, 237–241 Warburg micro syndrome and Martsolf syndrome, 253 western blotting, 263 posttranslational modifications, 236 REP. See Rab escort protein Reproductive life cycle, HPG axis critical biological rhythm, 72–73 estrous cycles, puberty, 72 GABA-GnRH connection adulthood, 97–107 early development, 92–94 puberty, 95–97 reproductive senescence, 107–108 glutamate/GABA interactions developmental transition, 110 GABAAR and GABABR, 110–112 neuron structural model, 109–110 OAA inhibitor, 111 POA examination, 111–112 P15 to P30 transition, 110–111 vesicular GABA transporter (VGAT), 91, 112 glutamate-GnRH connection adulthood, 83–86 early development, 78–79 puberty, 79–83 reproductive senescence, 86–90 GnRH–glutamate–GABA connection, 74–75 HPG axis, 71, 75 senescence age-related changes, receptor expression, 86–87 aging affects glutamate levels, 90 anteroventral peroventricular nucleus, 88 coexpression changes, 87–89 functional changes, 89 GABA-GnRH connection, 107–108

hypothalamic hormones measurement, 73–74 total reproductive failure, 73 Reproductive senescence, HPG axis age-related changes, receptor expression, 86–87 aging affects glutamate levels, 90 anteroventral peroventricular nucleus, 88 coexpression changes, 87–89 functional changes, 89 GABA-GnRH connection, 107–108 hypothalamic hormones measurement, 73–74 total reproductive failure, 73 Reticulons, 316 S Saccharomyces cerevisiae, 316 Scandentia, 309–310 Sexual maturation, 71–72 Smooth muscle contraction, h-CaD ERK and p38MAPK, 25–26 PAK phosphorylation, 26–27 phosphorylation, 25 SNARE proteins membrane fusion machinery, 135, 137 molecular/cellular research biochemistry, cytology, and forward genetics, 202 cytokinesis, 215 ER–Golgi trafficking, 206–207 forward genetic approaches, 201 secretion, 209–211 vacuolar and PVC /endosomal trafficking, 213–214 T Tip growth, higher order plant functions pollen tube growth, 218 root hair growth, 218–219 spatio-temporal regulator, 217 Trafficking pathways, 131–132 Trans-Golgi network (TGN), 131 Tubulo-reticular structure (TRS) autoimmune disease, 321 membrane symmetry, 280–301 Tupaia belangeri, 326 V Vacuolar and prevacuolar compartments (PVC)/ endosomal trafficking RAB18/RABC, 213 RAB5/RABF, 211–213 RAB7/RABG, 211 SNARE, 213–214 Vacuolar apical compartment (VAC), 146

350 Vesicular GABA transporter (VGAT), 91, 112 Vesicular trafficking biochemistry, cytology, and forward genetics RAB, 201–202 SNARE, 202 plant RAB and SNARE, initial identification, 200–201 post genome era, 203 transport pathway, RABs and SNAREs, 203–205 Virus-infected cells, 319–320 von Hippel–Lindau protein (VHL), 155

Index W Warburg micro syndrome, 253 Wiskott–Aldrich syndrome (WAS), 161–162 X Xenopus laevis, 309 Z Zebrafish system, in vivo model, 142 Zygnema sp., 303–304, 311, 326