Cutting Proteins within Lipid Bilayers: Rhomboid Structure and Mechanism

Molecular Cell

Review Cutting Proteins within Lipid Bilayers: Rhomboid Structure and Mechanism Marius K. Lemberg1,2 and Matthew Freeman1,* 1MRC

Laboratory of Molecular Biology, Hills Road, Cambridge CB2 0QH, UK address: Center for Molecular Biology, University of Heidelberg (ZMBH), Im Neuenheimer Feld 282, 69120 Heidelberg, Germany. *Correspondence: [email protected] DOI 10.1016/j.molcel.2007.12.003 2Present

Rhomboids were only discovered to be novel proteases in 2001, but progress on understanding this newest family of intramembrane proteases has been rapid. They are now the best characterized of these rather mysterious enzymes that cleave transmembrane domains within the lipid bilayer. In particular, the biochemical analysis of solubilized rhomboids and, most recently, a flurry of high-resolution crystal structures, have led to real insight into their enzymology. Long-standing questions about how it is possible for a water-requiring proteolytic reaction to occur in the lipid bilayer are now answered for the rhomboids. Intramembrane proteases, which control many medically important biological processes, have made the transition from rather heretical outsiders to novel enzymes that are becoming well understood. Introduction Over the last 10 years several new families of proteases have been identified that share the unexpected property of cleaving transmembrane (TM) domains of integral membrane proteins. Although enzymes catalyze all kinds of cellular reactions, the cleavage of peptide bonds in the plane of the lipid bilayer is unexpected, because proteolysis requires water, which is not abundant in a hydrophobic membrane. This has made the concept of intramembrane proteolysis slow to be fully accepted. Alternative suggestions to explain TM domain cleavage have included the presence of surface-exposed active sites, similar to the bacterial outer membrane protein OmpT (Vandeputte-Rutten et al., 2001), or the possibility that substrate TM domains are dislocated from the membrane before cleavage by soluble proteases. The first intramembrane protease to be discovered was the site 2 protease (S2P), the most famous is presenilin/g-secretase and its relatives, and the newest is the rhomboid family. But progress on rhomboids has been extremely rapid, and despite being the most recent intramembrane proteases to be discovered, they are now the family in which the mechanism of intramembrane proteolysis is best understood in atomic detail. Intramembrane proteases are found in all branches of life, and their functions are very diverse, including transcriptional control, cellular signaling, control of mitochondrial membrane remodeling, parasite invasion, and bacterial protein translocation (for reviews see Brown et al., 2000; Haass and Steiner, 2002; Urban and Freeman, 2002; Weihofen and Martoglio, 2003; Wolfe and Kopan, 2004). They have also been implicated in a wide range of human diseases, from Alzheimer’s to infection by pathogens including hepatitis C virus, M. tuberculosis, and the malaria parasite P. falciparum, making them potentially

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valuable new drug targets (Makinoshima and Glickman, 2005; Martoglio and Golde, 2003; O’Donnell et al., 2006; Selkoe and Schenk, 2003). Like the classical and well-understood soluble proteases, intramembrane proteases are classified according to the nucleophilic or water activating group that attacks the carbonyl group of a scissile peptide bond (Rawlings et al., 2006). The S2P family are metalloproteases (Akiyama et al., 2004; Duncan et al., 1998; Rawson et al., 1997); the GxGD family, the largest class, with up to seven members in humans including presenilin/g-secretase and signal peptide peptidase (SPP), are aspartyl proteases (De Strooper et al., 1998; Fluhrer et al., 2006; Friedmann et al., 2006; Narayanan et al., 2007; Weihofen et al., 2002; Wolfe et al., 1999); and rhomboids, the main focus here, are serine proteases, with five active members in humans (Lemberg and Freeman, 2007; Lemberg et al., 2005; Urban et al., 2001; Urban and Wolfe, 2005). The first rhomboid was identified as a Drosophila mutation with an abnormally pointed (rhomboid-shaped) head skeleton (Mayer and Nu¨sslein-Volhard, 1988). It was later found to be the primary regulator of EGF receptor (EGFR) signaling in Drosophila and to be the prototype of a widely conserved family of related polytopic membrane proteins (Wasserman et al., 2000). Their function as proteases was discovered when it was shown that Drosophila Rhomboid1 catalyzes the release of membrane-tethered growth factors, providing the trigger in the generation and secretion of the bioactive signal (Lee et al., 2001). Rhomboids are found in almost all species (Koonin et al., 2003; Lemberg and Freeman, 2007), with additional functions now known to range from control of mitochondrial membrane fusion (Herlan et al., 2003; McQuibban et al., 2003, 2006), to parasite invasion (Brossier et al., 2005; O’Donnell et al., 2006), to the activation of bacterial protein translocase crucial for

Molecular Cell

Review the generation of a peptidic quorum sensing signal (Stevenson et al., 2007). Recently the mitochondrial rhomboid has also been implicated in apoptosis (Cipolat et al., 2006). The biological functions of rhomboids have been reviewed extensively (Freeman, 2004; Pellegrini and Scorrano, 2007; Urban, 2006). Here, we focus instead on recent advances in understanding their mechanism and structures. Most significantly, a number of landmark papers have recently reported their high-resolution crystal structures—the first of any intramembrane protease. A very recent breakthrough paper has also reported the solution of the first crystal structure of an S2P family protease (Feng et al., 2007). All doubts that hydrolysis can occur in the hydrophobic environment of the membrane have now evaporated, and we have real mechanistic insights into how rhomboids and S2P work. Our goal is to describe the great strides in understanding rhomboids that have been made, and to highlight both the common ground and the areas of dispute, from which quite specific questions can now be framed. Biochemical Analysis of Rhomboids The initial evidence that rhomboid was a novel intramembrane protease depended on genetic and cell culture analysis. Specifically, the key observation was the demonstration that Drosophila Rhomboid-1 was able to cleave the growth factor Spitz (a Drosophila equivalent of mammalian TGFa) when they were coexpressed in COS-7 cells (Lee et al., 2001). Sequence conservation combined with mutational analysis and inhibitor studies led to the proposal that it was the first intramembrane serine protease, and that it probably used a serine-histidine-asparagine variant of the classical catalytic triad (Urban et al., 2001). Although these early studies demonstrated that rhomboids have the hallmarks of serine proteases, the cell-based assay was indirect and hampered a more detailed enzymological analysis. The putatively membrane-buried active site made developing an in vitro activity assay challenging, but three groups achieved this at about the same time (Lemberg et al., 2005; Maegawa et al., 2005; Urban and Wolfe, 2005). The details varied, but the principle was similar: bacterially expressed rhomboid was solubilized with detergent and mixed with an engineered protein based on a TM domain from a known rhomboid substrate. Activity could be detected in either crude extracts or purified fractions, and this provided the unambiguous proof that rhomboids are indeed proteases and that no essential cofactors are required. Cell-free analysis of rhomboid proteolysis implied that the enzyme uses a serine protease catalytic dyad of serine and histidine instead of the previously proposed triad (Lemberg et al., 2005); it also allowed accurate mapping of the substrate cleavage site and some basic biochemical properties of the enzyme to be explored. Although the solubilized enzymes have robust proteolytic activity, the enzyme works in vivo within a lipid bilayer, raising a question about the relevance of these assays. However,

they were validated by showing the same activity in intact membranes (Lemberg et al., 2005) and reconstituted proteoliposomes (Urban and Wolfe, 2005). It was also shown that lipid composition can affect rhomboid activity, at least in a cell-free system (Urban and Wolfe, 2005). In summary, these in vitro studies led to a refined mechanistic model with a high degree of confidence that rhomboids are in fact serine proteases but that, in a variation on a theme, they rely on a single histidine activating the serine nucleophile, instead of a classical catalytic triad. A Breakthrough: The Structure of a Mysterious Enzyme Despite these biochemical advances, solution of the central mysteries of rhomboid-catalyzed intramembrane proteolysis required an atomic resolution structure. In a landmark paper, Ha and colleagues published the first high-resolution structure of any intramembrane protease: the 2.1 A˚ crystallographic structure of the E. coli rhomboid GlpG (Wang et al., 2006). This was quickly followed by three other groups, using either molecular replacement techniques, which rely on the coordinates of the first paper (Lemieux et al., 2007; Wu et al., 2006), or using their own coordinates (Ben-Shem et al., 2007). Lemieux et al. (2007) solved the H. influenzae GlpG instead of the E. coli enzyme. To add to this flurry, two further structures have since been published by the Ha group (Wang and Ha, 2007; Wang et al., 2007). Importantly, although crystallized in different detergents and space groups, the architecture of GlpG from all these structures is very similar. The variations are relatively minor, but they are significant, and there are still uncertainties, which we discuss below. What do we learn? First, intramembrane proteolysis really does occur within the membrane. As shown in Figure 1, the rhomboid active site is located about 10 A˚ below the expected membrane surface. Note, however, that a very recent report suggests that the lipid bilayer might be distorted by the rhomboid TM helices to be abnormally thin, which would place the active site closer to the surface of the membrane (Wang et al., 2007). Second, we have the structural answer to the problem of how hydrolysis occurs in this environment: although nothing is yet known about substrate binding, the structure of the apoenzyme (in the absence of substrate or inhibitor) implies that proteolysis occurs in a water-filled indentation. The membrane-integral portion of GlpG is dominated by a bundle of six TM helices. Five of the six span the entire membrane; in the center of this bundle, the catalytic serine is positioned at the top of a shorter, tilted TM helix 4 (Figure 1). This arrangement thereby forms a water-filled cavity on the extracellular side of GlpG, with the catalytic serine at its base. In accordance with this architecture, the membrane-integral active site had been shown to be water accessible by biochemical crosslinking (Maegawa et al., 2007). The third feature agreed on by all rhomboid structures is the geometry of the active-site residues: rhomboids use a catalytic dyad between serine (S201 in E. coli GlpG) and histidine (H254) in TM helix 6 (Figure 2).

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Review Figure 1. Molecular Structure of the E. coli Rhomboid GlpG Ribbon diagram of E. coli GlpG, taken from coordinates of Wang et al. (2006) (PDB code 2IC8). See text for description of overall architecture. The loop connecting TM helices 1 and 2 (labeled L1, bottom panel, left) forms a novel protein fold extending sidewise into the plane of the membrane. For clarity, no secondary structure is indicated for the two non-TM helices of L1 in the front and back view (top panel). In order to show clearly the hydrophilic indentation and the proposed gate and cap structures, L1 and L5 are not shown in the surface representation (bottom panel, right). The white asterisk indicates the position of the catalytic center.

The conserved asparagine (N154) in TM helix 2, which initially had been suggested to participate in a chymotrypsin-like catalytic triad (Urban et al., 2001), is not in hydrogen bonding distance, and its function remains unclear. A prominent and unusual structural feature is the L1 loop, between TM helices 1 and 2, which extends sidewise

into the periplasmic leaflet of the lipid bilayer, with the highly conserved tryptophan-arginine motif ‘‘snorkeling’’ out of the hydrophobic membrane core (Wang et al., 2007), possibly providing a well-defined position in the membrane head-group region (Liang et al., 2005). The L5 loop (between TM helices 5 and 6) is also of interest, as there has been discussion that it might cap the active site in a closed state (see below). The major difference between the published structures is the position of TM helix 5 (Figure 3). In the first published structure, the active site is apparently shielded from all sides by helices, and substrate was speculated to enter between TM helices 1 and 3, with a gating mechanism dependent on L1. In other structures, TM helix 5 was tilted differently or was less ordered, suggesting potential substrate access from the opposite side of the molecule. We will discuss these differences and their implications below, but we wish to emphasize here that on most of the big questions, all structures agree, and that, in combination with the recent advances in the biochemistry, they begin to explain how these unusual proteases cleave peptide bonds within the membrane.

Figure 2. Rhomboid Active Site Highly conserved residues are shown in stick representation. The active-site structure illustrates the serine protease catalytic dyad (between S201 and H254 of E. coli GlpG). Conserved residues in TM helix 2 are not in hydrogen bonding distance and may fulfill other functions, such as substrate binding. Note that the tyrosine (Y205), which has been proposed to help stabilize the position of the catalytic histidine (H254), is not strictly conserved, so that in some rhomboids the histidine may be stabilized by other contacts. Ribbon representation as in Figure 1, back view.

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What Can We Infer about the Rhomboid Catalytic Mechanism? The hydrophilic cavity surrounding the rhomboid active site suggests an aqueous microenvironment suitable for a hydrolysis reaction to occur in the plane of the membrane. As in classical proteases, the catalytic serine

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Figure 3. Alternative Conformations for the Putative Gating Helix 5

Figure 4. Model for Rhomboid Structure Function Relationship

The GlpG structure reported by Wang et al. (2006) (shown in red; structure as in Figure 1) was superimposed on the altered conformation reported by Wu et al. (2006) (shown in yellow; PDB code 2NRF, molecule A). The outward bending of TM helix 5 (H5) may act as a gate, shielding the active site from the lipid environment in the closed state (red), and opening (yellow) to accommodate entering substrate (Wu et al., 2006).

A front view, in which the positions of highly conserved residues are indicated, is shown. For clarity, the putative lateral gate L1 is moved within the plane of the membrane. Red arrows indicate this and other suggested movements, potentially providing the gate for substrate entry and a surface cap structure (GlpG regions with apparent high structural plasticity are highlighted in pale blue). Two proposed substrate entry routes are indicated by green arrows (see text for details).

must be activated for its nucleophilic attack of the scissile peptide bond. Biochemical data combined with the recent structures suggest that this is mediated by the invariant histidine (H254) stripping away the proton from the serine (S201) (Lemberg et al., 2005; Wang et al., 2006). Typically, in classical serine proteases the histidine is in close proximity to a negatively charged aspartate, stabilizing the active conformation of the enzyme (Fersht, 1999; Hedstrom, 2002). For rhomboids, however, histidine alone appears to be sufficient to act as a general base, resulting in a serinehistidine catalytic dyad (Figure 2). In this noncanonical mechanism, the histidine may be stabilized by secondary contacts such as stacking against the p electrons of a tyrosine residue (Y205) (Baker et al., 2007; Wang et al., 2006). Another feature of serine proteases is a pocket stabilizing the negatively charged oxyanion caused by nucleophilic attack of the scissile carbonyl group. Typically this ‘‘oxyanion hole’’ is formed by two main-chain amino groups: for example, the serine and the first glycine of the GDSGG motif surrounding the active serine in chymotrypsin (Fersht, 1999). The GASG motif common in the rhomboid active site, combined with the requirement for the first glycine in functional assays, led to the suggestion of a related oxyanion hole in rhomboids (Lemberg et al., 2005; Urban et al., 2001, 2002). In the apoenzyme, however, only the backbone of the serine is in the right conformation to stabilize a putative oxyanion (Wang et al., 2006), making this unlikely without substantial conformation changes. Alternatively, the partial positively charged dipole of TM helix 4 (Lemieux et al., 2007) and conserved polar side chains nearby may contribute to stabilization of the negatively charged transition state. Various models involving the asparagine (N154) and/or the histidine (H150) (Ben-Shem et al., 2007; Lemieux et al., 2007; Wang and

Ha, 2007) are consistent with the influence of these residues in functional assays (Baker et al., 2007; Lemberg et al., 2005; Urban et al., 2001, 2002).

Substrate Entry A major challenge, and a point of some disagreement, is to explain how rhomboid substrates translocate from the lipid bilayer into the polar active site. In the absence of structures of enzyme/substrate complexes, any answers are necessarily speculative. Two different rhomboid substrate access models have been suggested, and three structural elements had been implicated in gating of the active site (Figure 4). In the first model to be proposed, substrate entry was envisioned to occur between TM helices 1 and 3, with the membrane-embedded L1 loop forming the putative lateral gate (Lemieux et al., 2007; Wang et al., 2006). This model implies substantial conformational changes, potentially rearranging the entire active-site conformation. This hypothesis is consistent with the apparent mobility in the crystal structure indicated by a high B factor, which suggests that this region is also flexible in the native state. A very recent structure has, however, led to the suggestion that the L1 loop may be quite rigid, casting doubt on its potential as a substrate gate (Wang et al., 2007). In the alternative access model, substrate may enter through a portal formed by TM helices 2 and 5 (Figure 4) (Ben-Shem et al., 2007; Wang and Ha, 2007; Wu et al., 2006). Two alternative gating mechanisms have been proposed for this access route. Wu et al. (2006) suggested that substrate entry is controlled by TM domain 5. This region shows the highest positional variation among all five structures (Figure 3), which is consistent with its possible

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Figure 5. Rhomboid Protease Consensus Conservation between rhomboids from multiple species that have been used in mutagenesis experiments is superimposed on the E. coli GlpG structure. Regions of highly conserved residues are indicated in red (>90% identity), orange is used for 80%–89% identity, and yellow is used for 50%–79% identity (see Lemberg and Freeman [2007] and references therein for details). The active-site consensus is GxSx in TM domain 4 (H4) and a single histidine (H) in TM domain 6 (H6). Small residues such as glycines (G) and alanine (A) in H4 and H6 allow tight helix packing that brings the catalytic serine and histidine in hydrogen bonding distance (Ben-Shem et al., 2007). The function of the conserved residues in TM domain 2 (H2) and the L1 loop is not yet clear. Note that the tryptophan-arginine motif (WR) of L1 is absent in mitochondrial PARL-type rhomboids but is strikingly conserved in the secretase-type and prokaryotic rhomboids. Residues implicated in the putative active-site gating mechanism centered around TM domain 5 and L5, such as F245 in E. coli GlpG, are not conserved (see text for details).

mobility (White, 2006). The hypothesis of substrate entry from this side is supported by an ordered phospholipid bound between TM helices 2 and 5 in the structure reported by Ben-Shem et al. (2007), indicating exposure of the active site to the lipid environment. This model is also consistent with recent enzymatic analysis of detergent-solubilized purified GlpG (Baker et al., 2007). The lack of obvious sequence conservation in this region, however, is surprising if it has such a critical functional role (Figure 5). Another potential gate for the gap between TM helices 2 and 5 is the L5 loop, which was proposed to cap the active site (Wang et al., 2006), segregating it from the extracellular environment (Figure 4). This region is metastable in the crystal packing, and in its more open conformation leads to the removal of a bulky phenylalanine side chain (F245), which, in the closed conformation, blocks access to the active site (Wang and Ha, 2007). Wu et al. (2006) observed a similar open conformation of the L5 loop in one of the two conformations they observed, potentially implicating it in the control of water supply. In summary, there are different models of gating and substrate access, but there appears to be growing sup-

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port for the idea that a space between TM helices 2 and 5 is perhaps the most plausible route, with the sequence variability of helix 5 possibly reflecting differences in rhomboid substrate specificity (Lieberman and Wolfe, 2007). We emphasize our earlier point, however, that this must remain speculative until enzyme/substrate structures are solved. The overall geometry of the rhomboid active site is solved, but again, in the absence of substrate in the crystal, the detailed mode of cleavage-site binding cannot be determined. The typical presence of helix-destabilizing residues near the cleavage site of substrate TM domains suggests that the substrate unfolds into the rhomboid active site (Akiyama and Maegawa, 2007; Urban and Freeman, 2003). This may be a prerequisite for the nucleophilic attack; indeed, proteases commonly bind their substrates in an extended b strand conformation (Tyndall et al., 2005). This model is consistent with the active-site geometry, which is predicted to be too narrow to accommodate the substrate in a helical conformation. Several attempts to model the docking of peptides into the GlpG active site have been made (Lemieux et al., 2007; Wang and Ha, 2007), leading to mechanistic speculations. For example, it has been suggested that the rhomboid catalytic serine attacks the scissile peptide bond from the si-face, opposite to most known serine proteases (Wang and Ha, 2007). Substrate Determinants Several studies show that rhomboids across evolution recognize related substrate features (Akiyama and Maegawa, 2007; Baker et al., 2006; Lemberg et al., 2005; Maegawa et al., 2005; Urban and Freeman, 2003; Urban et al., 2001; Urban and Wolfe, 2005). For example, rhomboids that localize in the eukaryotic secretory pathway cleave type I membrane proteins with helix-destabilizing residues like glycine near their cleavage site (Urban and Freeman, 2003). Consistent with this, a recent study showed a negative correlation between the proclivity of invariant residues to form stable TM helices and cleavage efficiency (Akiyama and Maegawa, 2007). Although substrate candidates have been identified based on such criteria (Lohi et al., 2004; Pascall and Brown, 2004; Urban and Freeman, 2003), no clear sequence consensus has been derived, and the molecular principles of rhomboid substrate selection remain poorly understood. For example, mammalian RHBDL2 can cleave the TM domain of thrombomodulin, even when the predicted helix-destabilizing features are mutated. In this case substrate determinants outside the TM domain were mapped (Lohi et al., 2004). Overall, we see an emerging picture where the propensity of a TM domain to be a rhomboid substrate is determined partially, but not fully, by helical instability: proteins with relatively unstable TM helices are good substrates for many rhomboids; but other TM domains can be substrates, if there are alternative interactions to help drive the enzyme-substrate binding. This may include recognition of motifs surrounding the scissile peptide bond, as is the case for classical soluble proteases; such primary

Molecular Cell

Review Figure 6. Active-Site and Substrate Topology Secretase and mitochondrial PARL-type rhomboids have active sites with opposite orientations. Comparative analysis of rhomboids showed variations of the basic six-TM-domain rhomboid core (Koonin et al., 2003; Lemberg and Freeman, 2007): most eukaryotic rhomboids, such as Drosophila Rhomboid-1, have an extra TM domain fused to the C terminus (indicated in red); mitochondrial PARL-type rhomboids, however, have an extra TM domain fused to the N terminus, thereby changing the orientation of the catalytic residues. The catalytic GASG and histidine of secretase rhomboids reside in TM helices 4 and 6, which both have out-to-in orientation (indicated by white arrowheads). In contrast, these catalytic motifs in PARLs are in the in-to-out helices 5 and 7. Intriguingly, there is a corresponding inversion of substrate orientation: PARL substrates have an Nin/Cout topology, but secretase rhomboids cleave type I membrane proteins (Nout/Cin). Upon Star-dependent transport to the Golgi compartment, cleavage by Drosophila Rhomboid-1 releases the N-terminal portion of the membrane-tethered growth factor Spitz, thereby allowing its secretion to trigger EGFR signaling (Lee et al., 2001). In contrast, the mitochondrial S. cerevisiae PARL (Pcp1/Rbd1) cleaves its substrate Mgm1 to release the C-terminal portion into the intermembrane space (IMS) (Herlan et al., 2003; McQuibban et al., 2003). Mgm1 processing is controlled by the ATP-dependent integration of the scissile TM domain via the TIM23 translocase (Herlan et al., 2004).

sequence preference has been observed for the E. coli rhomboid GlpG (Akiyama and Maegawa, 2007). The rhomboids studied so far all cleave their substrates at or close to the hydrophobic/hydrophilic boundary of the membrane—in most cases this is the N-terminal end of the TM domain, at the extracellular/luminal/periplasmic face of the membrane. The precise location of the scissile bond and how this is determined is unclear. GlpG was shown to cleave a serine-aspartate peptide bond located just N terminal of the TM domain of a model substrate, and even has some low activity against soluble casein (Maegawa et al., 2005). In all other cases, cleavage sites are within predicted TM helices (Baker et al., 2007; Herlan et al., 2004; Lemberg et al., 2005; Lohi et al., 2004; O’Donnell et al., 2006; Pascall and Brown, 2004; Urban et al., 2001). Thiol crosslinking in a single cysteine mutagenesis scan suggests that the GlpG cleave site is water accessible, potentially implying that it is exposed to the periplasm (Maegawa et al., 2007). Note, however, that this technique needs to be interpreted with care, as has been discussed by Grziwa et al. (2003). Overall, current evidence suggests that the substrate may be able to bend, so that its juxtamembrane region can, in some cases, enter the plane of the membrane in order to get access to the rhomboid active site. Rhomboid Active-Site Topology Rhomboids are a rare example of a membrane protein family in which the topology has evolved by gene fusion

adding a single TM domain to a conserved core. This leads to a number of different topological classes of rhomboids (Koonin et al., 2003; Lemberg and Freeman, 2007). Like most prokaryotic rhomboids, the E. coli rhomboid GlpG has the core six-TM-domain structure (Daley et al., 2005; Maegawa et al., 2005). Rhomboids in the second class, which are mainly found in the eukaryotic secretory pathway (termed secretase-type rhomboids) and some prokaryotes such as B. subtilis, have an extra TM domain fused to the C terminus (Figure 6) (Urban et al., 2001). Finally, in a third class, mitochondrial rhomboids, with human PARL as its most prominent member, all have an extra TM domain fused to the N terminus. Although the rhomboid active site is conserved, the addition of an Nterminal TM domain leads to the inversion of the active site (Figure 6) (Lemberg and Freeman, 2007). Intriguingly, the substrates of these mitochondrial rhomboids have a matching inversion, which is consistent with a widely held, but admittedly unproven, view that rhomboids— and indeed perhaps all intramembrane proteases—only cleave one orientation of substrate. An apparent exception to this idea is the recently reported cleavage by a secretase rhomboid of a type II membrane protein, the Drosophila transport chaperone Star (Tsruya et al., 2007). If Star cleavage is indeed directly catalyzed by rhomboid, it implies that rhomboid active sites can bind TM domains in either orientation. Enzymesubstrate interactions, stabilizing the transition state of

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Review proteolysis, are commonly very tight and occur in a regioand stereo-specific manner (Fersht, 1999; Milton et al., 1992; Roques et al., 1983). The scissile peptide bond and amino acid residues surrounding it are asymmetric, implying that for cleavage of both orientations with equal efficiency, the protease active-site determinants require a two-fold symmetry axis. This has been observed for retropepsin-type aspartyl proteases, which have a symmetric active site formed by a homodimer interface (Wlodawer and Gustchina, 2000). The question of whether rhomboids can cleave TM domains in both orientations remains open, but the observation by Tsruya et al. (2007) is intriguing, and the issue of the possible symmetry of the active site can now be systematically addressed with the biochemical and structural tools we have. Regulation of Rhomboids Cellular proteolysis is irreversible and directs changes ranging from protein degradation to precise signaling. Uncontrolled protease activity is therefore dangerous and is linked to many human diseases. Rhomboids differ from all the other families of intramembrane proteases by not requiring prior trimming of the substrate: rhomboids cleave intact membrane proteins. Without this regulated prior cleavage, how is rhomboid activity controlled? The full extent of potential regulation is unknown, but some principles have emerged in the last few years. In fact, most of the physiological control of rhomboids described so far involves regulating whether enzyme and substrate are in the same place at the same time; in contrast, almost nothing has been published about physiologically significant modulation of enzyme activity. The simplest example of control by access is transcriptional: Drosophila Rhomboid-1 is expressed in a precise and complex pattern that prefigures EGFR activity (reviewed in Freeman, 2004). This expression pattern is in turn determined by developmental signaling pathways that act on the rhomboid-1 enhancer (Nakamura et al., 2007; Sudarsan et al., 2002). The importance of this tight expression control is illustrated by the observation that, in most contexts, ectopic expression of rhomboid is sufficient to trigger EGFR signaling (because the membrane-tethered ligand and the receptor are widespread). The best-characterized mode of rhomboid regulation is cellular compartmentalization. Again, this was first discovered in Drosophila. The principal EGFR ligand, Spitz, is synthesized as a membrane-anchored proform that has to be cleaved by rhomboid for the extracellular domain to be secreted as a bioactive ligand (Figure 6). Rhomboid-1 is located in post-endoplasmic reticulum compartments, but Spitz is confined to the ER, thereby segregating it from the enzyme. A third membrane protein, Star, acts as a transport chaperone that escorts Spitz to the Rhomboid-1-containing compartment, thereby triggering its processing by Rhomboid-1 (Lee et al., 2001; Pascall et al., 2002; Tsruya et al., 2002). Extensive genetic analysis strongly implies that this regulated compartmentalization is the primary mode of controlling Spitz cleavage by Rhomboid-1 (Bang

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and Kintner, 2000; Guichard et al., 1999; Lee et al., 2001; Tsruya et al., 2007), but this does not imply that the same will be true in all cases. Nevertheless, there is evidence to suggest that rhomboid control by compartmentalization is not confined to Drosophila. In the apicomplexan parasite Toxoplasma gondii, rhomboid substrate adhesion proteins are segregated away from the active enzyme in a specific organelle, the microneme, until invasion occurs; they only meet each other during the invasion reaction, and the subsequent rhomboid-catalyzed cleavage may be an important process in host invasion (Brossier et al., 2005). An interesting variation of controlled substrate presentation is found in yeast mitochondria. The dynamin-like GTPase Mgm1 is a substrate of the PARL-type mitochondrial rhomboid Pcp1/Rbd1 (Figure 6) (Herlan et al., 2003; McQuibban et al., 2003). In this case, however, the primary Mgm1 TM domain is not the substrate. Instead, mitochondrial import machinery pulls Mgm1 further into mitochondrial matrix until a second potential TM domain, harboring the rhomboid cleavage site, becomes integrated into the inner mitochondrial membrane (Herlan et al., 2004). This second translocation requires ATP, thereby coupling Mgm1 processing with the mitochondrial bioenergetic state (Herlan et al., 2004). The balance between the membrane-anchored and the rhomboid-released isoforms of Mgm1 is crucial for the control of mitochondrial membrane dynamics in yeast (Herlan et al., 2003; McQuibban et al., 2003). A similar process occurs with yeast cytochrome c peroxidase (Ccp1), the second known substrate of Pcp1/Rbd1: the primary TM domain has to be dislocated by the m-AAA protease in order to allow rhomboid cleavage (Esser et al., 2002; Tatsuta et al., 2007). By analogy with other secretases and sheddases in the secretory pathway, it seems likely that rhomboid enzymatic activity itself is also directly regulated, and although there is no evidence yet to confirm this, there are some tantalizing observations. For example, it has been suggested that the activity of the mammalian mitochondrial rhomboid PARL may be regulated by phosphorylation and autoproteolytic processing in the N-terminal tail (Jeyaraju et al., 2006). Equally, some rhomboids have putative calcium-binding EF hands (Koonin et al., 2003) or a lipid-binding domain (Del Rio et al., 2007); these could represent regulatory sites. Even more speculatively, a class of highly conserved, catalytically inactive rhomboid homologs (termed iRhoms) could regulate rhomboid activity (Lemberg and Freeman, 2007). As the number of known biological and medically relevant functions for rhomboids increases, the importance of understanding their regulation grows, and it seems likely that this will be an area of intense focus in the next years. Similarities and Contrasts with Other Intramembrane Proteases The different families of intramembrane proteases are structurally and evolutionarily unrelated, and there is no reason to believe that they share a common mechanism. Nevertheless, all face the same fundamental problems in

Molecular Cell

Review Table 1. Summary Conclusions—A Rhomboid Primer Rhomboids are intramembrane serine proteases. Like other intramembrane proteases, they catalyze cleavage of transmembrane domains in many biological and medical contexts. The key active-site residues have been clearly identified by a combination of genetics, biochemistry, and structural biology: the active site depends on a novel catalytic dyad instead of the classical triad of other serine proteases. High-resolution crystal structures of rhomboids have recently been solved—the first for any intramembrane protease. Structures confirm that proteolysis occurs within the hydrocarbon region of the lipid bilayer, and functional studies showed that both cleavage products can be released from the membrane (see Figure 6). The mystery of how a hydrolytic reaction occurs in a lipid bilayer is solved by the discovery of a hydrophilic indentation in the extracellular face of the enzyme. Rhomboids have a conserved six-TM-domain core, but there are characteristic variations: most secretase rhomboids have an extra TM domain fused to the C terminus, while PARL-type rhomboids have an extra N-terminal TM domain. The structural consequences of this topology evolution remain to be addressed. More work is needed to solve current uncertainty about how the substrate enters the active site from the lipid bilayer. Little is yet known about the regulation of rhomboid enzyme activity: in vivo, there is significant regulation by compartmentalization of enzyme and substrate in the eukaryotic secretory pathway; in mitochondria, processing by rhomboid is linked to substrate topogenesis control. Rhomboids are now the best understood of all intramembrane proteases, and their analysis has revealed some principles that are applicable to all these enzymes. However, there is also evidence to suggest that each family may have distinct catalytic mechanisms.

recognizing their substrate, unfolding their TM helix, and performing the proteolysis reaction in the hydrophobic environment of the lipid bilayer. It therefore seems likely that understanding these processes in rhomboids will also provide some insight into the mechanism of S2P- and GxGD-type proteases. Intramembrane proteases cleave their substrates in different positions: rhomboids cleave near the luminal/extracellular side of the membrane, S2P near the cytosolic surface, and GxGD-type proteases commonly in the center (presenilin/g-secretase cleaves at multiple sites, including the S3 site close to the cytosolic side) (for review see Weihofen and Martoglio, 2003; Wolfe and Kopan, 2004). This implies fundamental differences in the active-site architecture, with GxGD-type protease possibly forming a channel-like structure. Consistent with this, a recent low-resolution structure of the active presenilin/g-secretase complex by single-particle reconstruction suggested the existence of a central chamber, which could provide the aqueous microenvironment required for proteolysis to occur (Lazarov et al., 2006). In further support, biochemical studies demonstrated the water accessibility of the active-site residues (Sato et al., 2006; Tolia et al., 2006). Intriguingly, a recent report implied a nonproteolytic function for presenilin, acting as an endoplasmic reticulum calcium leak channel (Tu et al., 2006). The recently reported structure of an S2P family protease (Feng et al., 2007) also shows an aqueous channel, this time originating from a membrane-buried catalytic zinc atom to the cytosolic side of the lipid bilayer. A common theme for all intramembrane proteolysis is cleavage of a membrane-embedded helical TM domain. As with rhomboids, for S2P and SPP there is a preference to cleave substrates harboring residues with a low helical

propensity (Akiyama et al., 2004; Lemberg and Martoglio, 2002; Ye et al., 2000), suggesting that these substrates must unwind into the enzyme active site to expose the scissile peptide bond (Lemberg and Martoglio, 2004). There appears to be no such requirement for presenilin/g-secretase (Lichtenthaler et al., 1999). This suggests that certain intramembrane proteases are more aggressive, having evolved the ability to cleave stable TM helices. Unlike all other intramembrane proteases, presenilin/g-secretase is a multiprotein complex with at least four different subunits (De Strooper, 2003; Wolfe, 2006), so it is possible that these may contribute to its ability to cleave stable helices. Recall also that cleavage of at least one rhomboid substrate has no obvious need for helix instability. In this case of RHBDL2 and thrombomodulin, it may not be that the enzyme is more aggressive generally, but that there is a particularly favored enzyme substrate interaction, apparently mediated by regions outside the TM domain (Lohi et al., 2004). The multisubunit nature of g-secretase also raises the question of whether rhomboids too might use cofactors. Genetics and biochemistry imply that there are no essential cofactors under experimental conditions, but current evidence does not exclude the possibility that, at least in some contexts in vivo, rhomboid cofactors may exist. Conclusions and Perspectives Although rhomboids were recognized as intramembrane proteases quite recently, and their physiological function in many organisms remains unknown, the recent progress in their biochemistry and structural biology now makes them the best understood family of these intriguing enzymes (see Table 1). The first atomic structures of intramembrane proteases have provided spectacular insights into their mechanisms and have removed the last doubts

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Review that hydrolysis of peptide bonds can take place in the membrane. Inevitably, the recent work has also posed new questions. Moreover, the rapid succession of publications about rhomboids over the last year has tended to emphasize differences and led to controversy. One of our goals has been to point out that, where there is disagreement, there is not yet enough information to allow a firm conclusion to be reached. We have also tried to emphasize the key open questions about rhomboid mechanisms that now need to be addressed. Among others, these include structural information about substrate binding, and physiological regulation of rhomboid activity. A detailed understanding of this class of enzymes will help to evaluate their physiological function and will accelerate the development of potent and specific rhomboid inhibitors. ACKNOWLEDGMENTS We thank Kvido Strisovsky, Vinothkumar Kutti Ragunath, and Colin Adrain for helpful comments and critical reading of the manuscript. M.K.L. was supported by a fellowship from the Swiss National Science Foundation. REFERENCES Akiyama, Y., and Maegawa, S. (2007). Sequence features of substrates required for cleavage by GlpG, an Escherichia coli rhomboid protease. Mol. Microbiol. 64, 1028–1037. Akiyama, Y., Kanehara, K., and Ito, K. (2004). RseP (YaeL), an Escherichia coli RIP protease, cleaves transmembrane sequences. EMBO J. 23, 4434–4442. Baker, R.P., Wijetilaka, R., and Urban, S. (2006). Two Plasmodium rhomboid proteases preferentially cleave different adhesins implicated in all invasive stages of malaria. PLoS Pathog. 2, e113. 10.1371/journal.ppat.0020113. Baker, R.P., Young, K., Feng, L., Shi, Y., and Urban, S. (2007). Enzymatic analysis of a rhomboid intramembrane protease implicates transmembrane helix 5 as the lateral substrate gate. Proc. Natl. Acad. Sci. USA 104, 8257–8262. Bang, A.G., and Kintner, C. (2000). Rhomboid and Star facilitate presentation and processing of the Drosophila TGF-alpha homolog Spitz. Genes Dev. 14, 177–186. Ben-Shem, A., Fass, D., and Bibi, E. (2007). Structural basis for intramembrane proteolysis by rhomboid serine proteases. Proc. Natl. Acad. Sci. USA 104, 462–466. Brossier, F., Jewett, T.J., Sibley, L.D., and Urban, S. (2005). A spatially localized rhomboid protease cleaves cell surface adhesins essential for invasion by Toxoplasma. Proc. Natl. Acad. Sci. USA 102, 4146–4151. Brown, M.S., Ye, J., Rawson, R.B., and Goldstein, J.L. (2000). Regulated intramembrane proteolysis: a control mechanism conserved from bacteria to humans. Cell 100, 391–398. Cipolat, S., Rudka, T., Hartmann, D., Costa, V., Serneels, L., Craessaerts, K., Metzger, K., Frezza, C., Annaert, W., D’Adamio, L., et al. (2006). Mitochondrial rhomboid PARL regulates cytochrome c release during apoptosis via OPA1-dependent cristae remodeling. Cell 126, 163–175. Daley, D.O., Rapp, M., Granseth, E., Melen, K., Drew, D., and von Heijne, G. (2005). Global topology analysis of the Escherichia coli inner membrane proteome. Science 308, 1321–1323. Del Rio, A., Dutta, K., Chavez, J., Ubarretxena-Belandia, I., and Ghose, R. (2007). Solution structure and dynamics of the N-terminal cytosolic domain of rhomboid intramembrane protease from Pseudomonas aer-

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uginosa: insights into a functional role in intramembrane proteolysis. J. Mol. Biol. 365, 109–122. De Strooper, B. (2003). Aph-1, Pen-2, and Nicastrin with Presenilin generate an active gamma-Secretase complex. Neuron 38, 9–12. De Strooper, B., Saftig, P., Craessaerts, K., Vanderstichele, H., Guhde, G., Annaert, W., Von Figura, K., and Van Leuven, F. (1998). Deficiency of presenilin-1 inhibits the normal cleavage of amyloid precursor protein. Nature 391, 387–390. Duncan, E.A., Dave, U.P., Sakai, J., Goldstein, J.L., and Brown, M.S. (1998). Second-site cleavage in sterol regulatory element-binding protein occurs at transmembrane junction as determined by cysteine panning. J. Biol. Chem. 273, 17801–17809. Esser, K., Tursun, B., Ingenhoven, M., Michaelis, G., and Pratje, E. (2002). A novel two-step mechanism for removal of a mitochondrial signal sequence involves the mAAA complex and the putative rhomboid protease Pcp1. J. Mol. Biol. 323, 835–843. Feng, L., Yan, H., Wu, Z., Yan, N., Wang, Z., Jeffrey, P.D., and Shi, Y. (2007). Structure of a site-2 protease family intramembrane metalloprotease. Science 318, 1608–1612. Fersht, A. (1999). Structure and Mechanism in Protein Science: A Guide to Enzyme Catalysis and Protein Folding (New York: W.H. Freeman). Fluhrer, R., Grammer, G., Israel, L., Condron, M.M., Haffner, C., Friedmann, E., Bohland, C., Imhof, A., Martoglio, B., Teplow, D.B., and Haass, C. (2006). A gamma-secretase-like intramembrane cleavage of TNFalpha by the GxGD aspartyl protease SPPL2b. Nat. Cell Biol. 8, 894–896. Freeman, M. (2004). Proteolysis within the membrane: rhomboids revealed. Nat. Rev. Mol. Cell Biol. 5, 188–197. Friedmann, E., Hauben, E., Maylandt, K., Schleeger, S., Vreugde, S., Lichtenthaler, S.F., Kuhn, P.H., Stauffer, D., Rovelli, G., and Martoglio, B. (2006). SPPL2a and SPPL2b promote intramembrane proteolysis of TNFalpha in activated dendritic cells to trigger IL-12 production. Nat. Cell Biol. 8, 843–848. Grziwa, B., Grimm, M.O., Masters, C.L., Beyreuther, K., Hartmann, T., and Lichtenthaler, S.F. (2003). The transmembrane domain of the amyloid precursor protein in microsomal membranes is on both sides shorter than predicted. J. Biol. Chem. 278, 6803–6808. Guichard, A., Biehs, B., Sturtevant, M.A., Wickline, L., Chacko, J., Howard, K., and Bier, E. (1999). rhomboid and Star interact synergistically to promote EGFR/MAPK signaling during Drosophila wing vein development. Development 126, 2663–2676. Haass, C., and Steiner, H. (2002). Alzheimer disease g-secretase: a complex story of GxGD-type presenilin proteases. Trends Cell Biol. 12, 556–562. Hedstrom, L. (2002). Serine protease mechanism and specificity. Chem. Rev. 102, 4501–4524. Herlan, M., Vogel, F., Bornhovd, C., Neupert, W., and Reichert, A.S. (2003). Processing of Mgm1 by the Rhomboid-type protease Pcp1 is required for maintenance of mitochondrial morphology and of mitochondrial DNA. J. Biol. Chem. 278, 27781–27788. Herlan, M., Bornhovd, C., Hell, K., Neupert, W., and Reichert, A.S. (2004). Alternative topogenesis of Mgm1 and mitochondrial morphology depend on ATP and a functional import motor. J. Cell Biol. 165, 167–173. Jeyaraju, D.V., Xu, L., Letellier, M.C., Bandaru, S., Zunino, R., Berg, E.A., McBride, H.M., and Pellegrini, L. (2006). Phosphorylation and cleavage of presenilin-associated rhomboid-like protein (PARL) promotes changes in mitochondrial morphology. Proc. Natl. Acad. Sci. USA 103, 18562–18567. Koonin, E.V., Makarova, K.S., Rogozin, I.B., Davidovic, L., Letellier, M.C., and Pellegrini, L. (2003). The rhomboids: a nearly ubiquitous family of intramembrane serine proteases that probably evolved by multiple ancient horizontal gene transfers. Genome Biol. 4, R19.

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Review Lazarov, V.K., Fraering, P.C., Ye, W., Wolfe, M.S., Selkoe, D.J., and Li, H. (2006). Electron microscopic structure of purified, active gammasecretase reveals an aqueous intramembrane chamber and two pores. Proc. Natl. Acad. Sci. USA 103, 6889–6894.

Nakamura, Y., Kagesawa, T., Nishikawa, M., Hayashi, Y., Kobayashi, S., Niimi, T., and Matsuno, K. (2007). Soma-dependent modulations contribute to divergence of rhomboid expression during evolution of Drosophila eggshell morphology. Development 134, 1529–1537.

Lee, J.R., Urban, S., Garvey, C.F., and Freeman, M. (2001). Regulated intracellular ligand transport and proteolysis control EGF signal activation in Drosophila. Cell 107, 161–171.

Narayanan, S., Sato, T., and Wolfe, M.S. (2007). A C-terminal domain of signal peptide peptidase defines a functional domain for intramembrane aspartic protease catalysis. J. Biol. Chem. 282, 20172–20179.

Lemberg, M.K., and Freeman, M. (2007). Functional and evolutionary implications of enhanced genomic analysis of rhomboid intramembrane proteases. Genome Res. 17, 1634–1646.

O’Donnell, R.A., Hackett, F., Howell, S.A., Treeck, M., Struck, N., Krnajski, Z., Withers-Martinez, C., Gilberger, T.W., and Blackman, M.J. (2006). Intramembrane proteolysis mediates shedding of a key adhesin during erythrocyte invasion by the malaria parasite. J. Cell Biol. 174, 1023–1033.

Lemberg, M.K., and Martoglio, B. (2002). Requirements for signal peptide peptidase-catalyzed intramembrane proteolysis. Mol. Cell 10, 735–744. Lemberg, M.K., and Martoglio, B. (2004). On the mechanism of SPPcatalysed intramembrane proteolysis; conformational control of peptide bond hydrolysis in the plane of the membrane. FEBS Lett. 564, 213–218. Lemberg, M.K., Menendez, J., Misik, A., Garcia, M., Koth, C.M., and Freeman, M. (2005). Mechanism of intramembrane proteolysis investigated with purified rhomboid proteases. EMBO J. 24, 464–472. Lemieux, M.J., Fischer, S.J., Cherney, M.M., Bateman, K.S., and James, M.N. (2007). The crystal structure of the rhomboid peptidase from Haemophilus influenzae provides insight into intramembrane proteolysis. Proc. Natl. Acad. Sci. USA 104, 750–754. Liang, J., Adamian, L., and Jackups, R., Jr. (2005). The membrane-water interface region of membrane proteins: structural bias and the antisnorkeling effect. Trends Biochem. Sci. 30, 355–357. Lichtenthaler, S.F., Wang, R., Grimm, H., Uljon, S.N., Masters, C.L., and Beyreuther, K. (1999). Mechanism of the cleavage specificity of Alzheimer’s disease g-secretase identified by phenylalanine-scanning mutagenesis of the transmembrane domain of the amyloid precursor protein. Proc. Natl. Acad. Sci. USA 96, 3053–3058. Lieberman, R.L., and Wolfe, M.S. (2007). Membrane-embedded protease poses for photoshoot. Proc. Natl. Acad. Sci. USA 104, 401–402. Lohi, O., Urban, S., and Freeman, M. (2004). Diverse substrate recognition mechanisms for rhomboids: thrombomodulin is cleaved by Mammalian rhomboids. Curr. Biol. 14, 236–241. Maegawa, S., Ito, K., and Akiyama, Y. (2005). Proteolytic action of GlpG, a rhomboid protease in the Escherichia coli cytoplasmic membrane. Biochemistry 44, 13543–13552. Maegawa, S., Koide, K., Ito, K., and Akiyama, Y. (2007). The intramembrane active site of GlpG, an E. coli rhomboid protease, is accessible to water and hydrolyses an extramembrane peptide bond of substrates. Mol. Microbiol. 64, 435–447. Makinoshima, H., and Glickman, M.S. (2005). Regulation of Mycobacterium tuberculosis cell envelope composition and virulence by intramembrane proteolysis. Nature 436, 406–409. Martoglio, B., and Golde, T.E. (2003). Intramembrane-cleaving aspartic proteases and disease: presenilins, signal peptide peptidase and their homologs. Hum. Mol. Genet. 12, R201–R206. Mayer, U., and Nu¨sslein-Volhard, C. (1988). A group of genes required for pattern formation in the ventral ectoderm of the Drosophila embryo. Genes Dev. 2, 1496–1511. McQuibban, G.A., Saurya, S., and Freeman, M. (2003). Mitochondrial membrane remodelling regulated by a conserved rhomboid protease. Nature 423, 537–541. McQuibban, G.A., Lee, J.R., Zheng, L., Juusola, M., and Freeman, M. (2006). Normal mitochondrial dynamics requires rhomboid-7 and affects Drosophila lifespan and neuronal function. Curr. Biol. 16, 982–989. Milton, R.C., Milton, S.C., and Kent, S.B. (1992). Total chemical synthesis of a D-enzyme: the enantiomers of HIV-1 protease show reciprocal chiral substrate specificity. Science 256, 1445–1448.

Pascall, J.C., and Brown, K.D. (2004). Intramembrane cleavage of ephrinB3 by the human rhomboid family protease, RHBDL2. Biochem. Biophys. Res. Commun. 317, 244–252. Pascall, J.C., Luck, J.E., and Brown, K.D. (2002). Expression in mammalian cell cultures reveals interdependent, but distinct, functions for Star and Rhomboid proteins in the processing of the Drosophila transforming-growth-factor-alpha homologue Spitz. Biochem. J. 363, 347–352. Pellegrini, L., and Scorrano, L. (2007). A cut short to death: Parl and Opa1 in the regulation of mitochondrial morphology and apoptosis. Cell Death Differ. 14, 1275–1284. Rawlings, N.D., Morton, F.R., and Barrett, A.J. (2006). MEROPS: the peptidase database. Nucleic Acids Res. 34, D270–D272. Rawson, R.B., Zelenski, N.G., Nijhawan, D., Ye, J., Sakai, J., Hasan, M.T., Chang, T.Y., Brown, M.S., and Goldstein, J.L. (1997). Complementation cloning of S2P, a gene encoding a putative metalloprotease required for intramembrane cleavage of SREBPs. Mol. Cell 1, 47–57. Roques, B.P., Lucas-Soroca, E., Chaillet, P., Costentin, J., and Fournie-Zaluski, M.C. (1983). Complete differentiation between enkephalinase and angiotensin-converting enzyme inhibition by retro-thiorphan. Proc. Natl. Acad. Sci. USA 80, 3178–3182. Sato, C., Morohashi, Y., Tomita, T., and Iwatsubo, T. (2006). Structure of the catalytic pore of gamma-secretase probed by the accessibility of substituted cysteines. J. Neurosci. 26, 12081–12088. Selkoe, D.J., and Schenk, D. (2003). Alzheimer’s disease: molecular understanding predicts amyloid-based therapeutics. Annu. Rev. Pharmacol. Toxicol. 43, 545–584. Stevenson, L.G., Strisovsky, K., Clemmer, K.M., Bhatt, S., Freeman, M., and Rather, P.N. (2007). Rhomboid protease AarA mediates quorum-sensing in Providencia stuartii by activating TatA of the twin-arginine translocase. Proc. Natl. Acad. Sci. USA 104, 1003–1008. Sudarsan, V., Pasalodos-Sanchez, S., Wan, S., Gampel, A., and Skaer, H. (2002). A genetic hierarchy establishes mitogenic signalling and mitotic competence in the renal tubules of Drosophila. Development 129, 935–944. Tatsuta, T., Augustin, S., Nolden, M., Friedrichs, B., and Langer, T. (2007). m-AAA protease-driven membrane dislocation allows intramembrane cleavage by rhomboid in mitochondria. EMBO J. 26, 325–335. Tolia, A., Chavez-Gutierrez, L., and De Strooper, B. (2006). Contribution of presenilin transmembrane domains 6 and 7 to a water-containing cavity in the gamma-secretase complex. J. Biol. Chem. 281, 27633–27642. Tsruya, R., Schlesinger, A., Reich, A., Gabay, L., Sapir, A., and Shilo, B.Z. (2002). Intracellular trafficking by Star regulates cleavage of the Drosophila EGF receptor ligand Spitz. Genes Dev. 16, 222–234. Tsruya, R., Wojtalla, A., Carmon, S., Yogev, S., Reich, A., Bibi, E., Merdes, G., Schejter, E., and Shilo, B.Z. (2007). Rhomboid cleaves Star to regulate the levels of secreted Spitz. EMBO J. 26, 1211–1220. Tu, H., Nelson, O., Bezprozvanny, A., Wang, Z., Lee, S.F., Hao, Y.H., Serneels, L., De Strooper, B., Yu, G., and Bezprozvanny, I. (2006).

Molecular Cell 28, December 28, 2007 ª2007 Elsevier Inc. 939

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Review Presenilins form ER Ca2+ leak channels, a function disrupted by familial Alzheimer’s disease-linked mutations. Cell 126, 981–993. Tyndall, J.D., Nall, T., and Fairlie, D.P. (2005). Proteases universally recognize beta strands in their active sites. Chem. Rev. 105, 973–999. Urban, S. (2006). Rhomboid proteins: conserved membrane proteases with divergent biological functions. Genes Dev. 20, 3054–3068. Urban, S., and Freeman, M. (2002). Intramembrane proteolysis controls diverse signalling pathways throughout evolution. Curr. Opin. Genet. Dev. 12, 512–518. Urban, S., and Freeman, M. (2003). Substrate specificity of rhomboid intramembrane proteases is governed by helix-breaking residues in the substrate transmembrane domain. Mol. Cell 11, 1425–1434.

Wasserman, J.D., Urban, S., and Freeman, M. (2000). A family of rhomboid-like genes: Drosophila rhomboid-1 and roughoid/rhomboid-3 cooperate to activate EGF receptor signaling. Genes Dev. 14, 1651–1663. Weihofen, A., and Martoglio, B. (2003). Intramembrane-cleaving proteases: controlled liberation of proteins and bioactive peptides. Trends Cell Biol. 13, 71–78. Weihofen, A., Binns, K., Lemberg, M.K., Ashman, K., and Martoglio, B. (2002). Identification of signal peptide peptidase, a presenilin-type aspartic protease. Science 296, 2215–2218. White, S.H. (2006). Rhomboid intramembrane protease structures galore! Nat. Struct. Mol. Biol. 13, 1049–1051.

Urban, S., and Wolfe, M.S. (2005). Reconstitution of intramembrane proteolysis in vitro reveals that pure rhomboid is sufficient for catalysis and specificity. Proc. Natl. Acad. Sci. USA 102, 1883–1888.

Wlodawer, A., and Gustchina, A. (2000). Structural and biochemical studies of retroviral proteases. Biochim. Biophys. Acta 1477, 16–34.

Urban, S., Lee, J.R., and Freeman, M. (2001). Drosophila rhomboid-1 defines a family of putative intramembrane serine proteases. Cell 107, 173–182.

Wolfe, M.S. (2006). The gamma-secretase complex: membrane-embedded proteolytic ensemble. Biochemistry 45, 7931–7939.

Urban, S., Schlieper, D., and Freeman, M. (2002). Conservation of intramembrane proteolytic activity and substrate specificity in prokaryotic and eukaryotic rhomboids. Curr. Biol. 12, 1507–1512. Vandeputte-Rutten, L., Kramer, R.A., Kroon, J., Dekker, N., Egmond, M.R., and Gros, P. (2001). Crystal structure of the outer membrane protease OmpT from Escherichia coli suggests a novel catalytic site. EMBO J. 20, 5033–5039. Wang, Y., and Ha, Y. (2007). Open-cap conformation of intramembrane protease GlpG. Proc. Natl. Acad. Sci. USA 104, 2098–2102. Wang, Y., Zhang, Y., and Ha, Y. (2006). Crystal structure of a rhomboid family intramembrane protease. Nature 444, 179–180. Wang, Y., Maegawa, S., Akiyama, Y., and Ha, Y. (2007). The role of L1 loop in the mechanism of rhomboid intramembrane protease GlpG. J. Mol. Biol. 374, 1104–1113.

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Wolfe, M.S., and Kopan, R. (2004). Intramembrane proteolysis: theme and variations. Science 305, 1119–1123. Wolfe, M.S., Xia, W., Moore, C.L., Leatherwood, D.D., Ostaszewski, B., Rahmati, T., Donkor, I.O., and Selkoe, D.J. (1999). Peptidomimetic probes and molecular modeling suggest that Alzheimer’s gamma-secretase is an intramembrane-cleaving aspartyl protease. Biochemistry 38, 4720–4727. Wu, Z., Yan, N., Feng, L., Oberstein, A., Yan, H., Baker, R.P., Gu, L., Jeffrey, P.D., Urban, S., and Shi, Y. (2006). Structural analysis of a rhomboid family intramembrane protease reveals a gating mechanism for substrate entry. Nat. Struct. Mol. Biol. 13, 1084–1091. Ye, J., Dave, U.P., Grishin, N.V., Goldstein, J.L., and Brown, M.S. (2000). Asparagine-proline sequence within membrane-spanning segment of SREBP triggers intramembrane cleavage by site-2 protease. Proc. Natl. Acad. Sci. USA 97, 5123–5128.