Article |
Address correspondence to Dr. Keith Joiner, LCI 808, Infectious Diseases Section, Department of Internal Medicine, Yale University School of Medicine, 333 Cedar St., New Haven, CT 06520. Tel.: (203) 785-4140. Fax: (203) 785-3864. E-mail: keith.joiner{at}yale.edu; sinai{at}pop.uky.edu
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Abstract |
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Key Words: Toxoplasma; ROP2; parasitophorous vacuole membrane; mitochondria; endoplasmic reticulum
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Introduction |
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The T. gondii PVM exhibits a remarkable association with host mitochondria and ER (DeMelo et al., 1992; Sinai et al., 1997) (see Fig. 1 A). We have previously termed this phenomenon PVMorganelle association (Sinai et al., 1997). Association of host organelles with the vacuolar membranes surrounding intracellular pathogens is a feature restricted to organisms that either never enter the endocytic cascade, or that exit it soon after entry (Sinai and Joiner, 1997). Both Legionella pneumophila (Swanson and Isberg, 1995) and Brucella abortus (Pizarro-Cerda et al., 1998) replicate in a compartment associated with the ER. In L. pneumophilainfected cells, a bacterially encoded machinery with homology to bacterial DNA conjugation systems is required for establishment and maintenance of the ER-associated replicative phagosome (Purcell and Shuman, 1998; Vogel et al., 1998). However, the substrates potentially being exported by this machinery remain elusive (Purcell and Shuman, 1998; Vogel et al., 1998). Even less well understood is the molecular basis for mitochondrial association observed with the inclusion (vacuole) membrane housing certain strains of Chlamydia psittaci (Matsumoto et al., 1991; Sinai and Joiner, 1997). Although host cytoplasmexposed proteins of chlamydial origin have been identified in the inclusion membrane, no link to mitochondrial association has been made (Rockey et al., 1997; Bannantine et al., 1998). What is clear is that both mitochondrial association by C. psittaci (Matsumoto et al., 1991) and ER association by L. pneumophila (Swanson and Isberg, 1995) and B. abortus (Pizarro-Cerda et al., 1998) are important in the establishment of the replication-permissive compartment, and likely involved in nutrient acquisition (Sinai and Joiner, 1997).
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Proteins modifying the T. gondii PVM originate in the excretory/secretory organelles of the parasite (Sinai and Joiner, 1997; Lingelbach and Joiner, 1998). These include club-shaped organelles called rhoptries (Dubremetz et al., 1998) (see Fig. 1 A), discharged concomitant with parasite invasion (Dubremetz et al., 1993; Carruthers and Sibley, 1997), and dense granules (see Fig. 1 A) that release their cargo throughout the intracellular residence of the parasite (Carruthers and Sibley, 1997). Morphometric data indicate that PVMorganelle association is established early in infection and does not increase with the time of intracellular residence, suggesting rhoptry involvement (Sinai et al., 1997). PVMorganelle association is poorly understood at the molecular and biochemical levels (Sinai and Joiner, 1997). The quest for the molecular basis of PVMorganelle association is further confounded by the fact that the physical nature of the interaction (i.e., Is it a proteinprotein, proteinlipid, lipidlipid, or other interaction?) is not known.
In this study, we describe the molecular mechanism by which the association between the T. gondii PVM and host cell organelles is established. The general mechanism reported here is one in which a PVM-anchored protein tethers host organelle membranes by inserting into them, promoting a stable association. This insight opens the way to defining the role of PVMorganelle association in parasite biology, and presents a mechanism by which organelle systems such as mitochondria and ER may interact in eukaryotes in general.
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Results |
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The kinetics for the establishment of PVMmitochondrial association, examined immediately after parasite invasion, were used to discriminate between the involvement of a rhoptry- or dense granulederived factor. Within 1 min of infection, 36% of intracellular T. gondii vacuoles had at least one mitochondrial profile associated with the PVM (Fig. 1 B, arrowheads). Notably, all of the staining for the PVM-localizing, dense granule marker GRA3 (Bermudes et al., 1994) was still retained within the parasite (Fig 1 B). The extent of PVMmitochondrial association increased to 68 and 74% of vacuoles at 5 and 10 min postinfection, respectively (Fig. 1 B). By these time points, dense granule exocytosis is apparent (Carruthers and Sibley, 1997), resulting in GRA3 release into the vacuolar space and the PVM (Fig. 1 B). Therefore, the kinetics of PVMmitochondrial association correlate better with rhoptry than with dense granule exocytosis (Fig. 1 B), as previously suggested (Sinai et al., 1997). This finding prompted us to examine ROP2, the founding member of a family of proteins known to be in the PVM (Beckers et al., 1994), as a potential mediator of PVMmitochondrial association.
The NH2-terminal domain of mature ROP2 is exposed to the host cell cytoplasm
In the infected cell, members of the ROP2 family are predominant antigens of the PVM where they are exposed to the host cell cytoplasm (Beckers et al., 1994). This earlier observation was in part based on labeling of the PVM with an antiserum against the rhoptry/dense granule (R/DG) fraction of the parasite, after selective permeabilization of infected host cells (Beckers et al., 1994). We sought to identify the domain within ROP2 containing the epitope(s) recognized by the R/DG antiserum.
The ROP2 protein is processed within the secretory pathway of the parasite en route to the rhoptry (Fig. 2 A) (Sadak et al., 1988; Leriche and Dubremetz, 1991). Although the precise processing site is unclear, NH2-terminal sequencing of an ROP2 fragment indicates that aa 98 is at or close to the processing site (Dubremetz, J.-F., personal communication) (Fig. 2 A). Therefore, we divided ROP2 into an NH2-terminal domain (aa 98465) and a COOH-terminal domain (aa 466561) around its predicted transmembrane region (Fig. 2 A). These domains were synthesized in vitro as 35S-Metlabeled substrates, and the ability of the R/DG antiserum to immunoprecipitate them was determined. The R/DG antiserum selectively immunoprecipitated the NH2-terminal domain of mature ROP2 (aa 98465; Fig. 2 B), but not a chimera of bacterial alkaline phosphatase (BAP) with the ROP2 transmembrane domain and COOH terminus (aa 466561) (Fig. 2 B). This observation indicates that the NH2-terminal domain of PVM-localized ROP2 is exposed to the host cytoplasm (RO2hc; aa 98465).
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Mitochondrial matrix targeting signals direct translocation into the matrix in a vectorial fashion led by the NH2 terminus (Neupert, 1997). The directionality of ROP2hc translocation was examined by exploiting the observation that the epitope recognized by the mAbT34A7 is included in the putative targeting signal (aa 98127) (unpublished data). mAb T34A7 immunoprecipitated full-length ROP2hc (Fig. 3 B, lane 7), and more significantly the 17-kD protease-protected fragment (Fig. 3 B, lane 8), from the mitochondrial pellet fractions in both the absence of and after protease treatment.
In all experiments, the import competence of the mitochondria was confirmed using human ornithine transcarbamylase (OTC) as a control (Horwich et al., 1986). OTC was correctly imported and processed from the full-length precursor protein (Fig. 3 C, p, lanes 13 and 14) to the mature form (Fig. 3 C, m, lanes 13 and 15) as a consequence of cleavage of the NH2-terminal targeting signal in the matrix (Horwich et al., 1986).
Because the processing site within ROP2 has not been precisely mapped and may be somewhat upstream of aa 98 (Fig. 2 A), we examined the interaction of a polypeptide identical to ROP2hc but originating at aa 80 (ROP2hc80) (Figs. 2 A and 3 A). Upon interaction with mitochondria, ROP2hc80 behaved exactly like ROP2hc, binding to the organelle pellet (Fig. 3 C, lane 5) and revealing a larger (19 kD) protease-resistant fragment (Fig. 3 C, lane 7, arrowhead) consistent with the increase in molecular weight imparted by the 18 aa from aa 8098 (Fig. 3 A). Because both ROP2hc and ROP2hc80 include the putative targeting signal (aa 98127), we sought to determine its contribution to translocation across the MOM in vitro.
The role of the putative targeting signal (aa 98127) in interaction with mitochondria was tested by deleting it to generate 98127ROP2hc. In the in vitro import assay,
98127ROP2hc still bound to mitochondria (Fig. 3 C, lane 9), but no protected fragment was observed after protease treatment (Fig. 3 C, lane 11). Together, these data indicate that translocation of the host cytoplasmexposed domain of ROP2 into the MOM in vitro is not critically dependent on the precise processing site, as long as the putative targeting signal (aa 98127) is present.
ROP2hc does not target the mitochondrial matrix
In light of the potential mitochondrial matrix targeting signal (aa 98127) in ROP2hc, we examined the effects of treatments known to block matrix import. We examined the consequences of temperature (0°C), ATP depletion (apyrase), dissipation of the membrane potential across the inner membrane (carboxyl cyanide m-chlorophenylhydrazone [CCCP]), and the requirement for trypsin- sensitive receptors on the mitochondrial surface such as TOM20 (reviewed in Neupert, 1997), on ROP2hc translocation. In addition, the ability of anti-TOM20 antibodies to block import was tested. In control experiments, the import and processing of the matrix-targeted protein OTC was blocked at 0°C (Fig. 4 A, OTC, lanes 58), and was significantly inhibited by treatment with apyrase (Fig. 4 A, OTC, lanes 912) or CCCP (Fig. 4 A, OTC, lanes 1316). As predicted, the mAb T34A7 against aa 98127 of ROP2hc, did not affect either the import or processing of OTC (Fig. 4 A, OTC, lanes 1720). In contrast, pretreatment of mitochondria with a chicken anti-TOM20 antibody completely blocked both the import and processing of OTC (Fig. 4 A, OTC, lanes 2124), as described previously (Goping et al., 1995).
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In contrast, the importance of the NH2-terminal signal of ROP2hc (aa 98127) in translocation is further supported by the lack of the 17-kD protease-protected fragment when the monoclonal T34A7 was added (Fig. 4 A, ROP2hc, lane 19, arrowhead). Notably, inhibition by T34A7 was observed only if the ROP2hc containing reticulocyte lysate was preincubated with the antibody before addition to mitochondria (unpublished data). In contrast, pretreatment of mitochondria with anti-TOM20 antibodies failed to block the generation of the protease-protected fragment (Fig. 4 A, ROP2hc, lane 27). The experiment is presented with an independent "no treatment" control (Fig. 4 A, ROP2hc, lanes 2124), as it was not performed at the time the other treatments were.
ROP2hc translocation does not require a trypsin-sensitive receptor
The role of trypsin-sensitive receptors in addition to TOM20 on targeting of imported proteins to the matrix and other mitochondrial compartments is well documented (Neupert, 1997). As reported previously (Argan et al., 1983), trypsin pretreatment of mitochondria completely blocked the import and processing of OTC (Fig. 4 B, lanes 1518). In contrast, the translocation of ROP2hc across the MOM of trypsin-pretreated mitochondria was not inhibited, resulting in a slightly smaller (15 kD) protected fragment (Fig. 4 B, lane 8, arrowhead), identical to that observed with nontreated mitochondria (Fig. 4 B, lane 4, arrowhead). Thus, despite possessing features of a matrix targeting signal, ROP2hc, in contrast to most imported mitochondrial proteins, translocates across the MOM without a requirement for a trypsin-sensitive receptor, supporting the results with the anti-TOM20 antibody (Fig. 4 A)
ROP2hc contains both mitochondrial- and ER-targeting domains
Next, we tested the targeting of ROP2hc in vivo. The level of expression of ROP2hc and ROP2hc80 (unpublished data) had a significant effect on the localization of the protein. Relatively low levels of ROP2hc resulted in localization to rod-shaped and punctate structures reminiscent of mitochondria (Fig. 5, A and C
, yellow arrowheads). These structures were confirmed to be mitochondria based on colocalization with the mitochondrial marker cytochrome c oxidase subunit III (COXIII) (Fig. 5, B and C, yellow arrowheads). Colocalization with mitochondria was confirmed with antibody to cytochrome c oxidase subunit 1 (COXI) (unpublished data), hamster MOM (unpublished data), and the mitochondrion-specific, membrane potentialsensitive dye MitoTracker (Molecular Probes) (Fig. 5, EG).
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We sought to determine whether the nonmitochondrial localization of ROP2hc corresponded to ER. ROP2hc (Fig. 5 H) exhibited some colocalization with the ER marker calnexin (Fig. 5 I). These appear as a yellow signal concentrated in the perinuclear area of the transfected cell in the merged image (Fig. 5 J). The red signal in the merged image (Fig. 5 J) likely corresponds to mitochondria. This observation suggests that ROP2hc localizes to both mitochondria and ER after expression in vivo.
Next, we examined whether deletion of the NH2-terminal 30 aa required for in vitro mitochondrial import played a role in ROP2hc localization in vivo. After expression of a 98127ROP2hcGFP chimera, a distinct pattern of localization was observed. Unlike ROP2hc,
98127ROP2hcGFP (Fig. 6, A, C, D, and F
, green arrowheads) did not target significantly to mitochondria, as detected with either MitoTracker (Fig. 6, B and C, red arrowhead) or anti-COXI antibody (Fig. 6, E and F, red arrowheads). Some colocalizing was observed (Fig. 6, DF, bottom cell, yellow arrowhead) and is likely due to the limitations in the resolving power of confocal microscopy. Such patches of apparent colocalization were only found where a high concentration of both signals was present (Fig. 6, D and E). Notably, expression of
98127ROP2hcGFP did not lead to the clumping and other changes in organelle organization (Fig. 6, B, C, E, and F) observed with ROP2hc (Fig. 5, B, C, E, and F).
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The contribution of the 30-aa putative mitochondrial targeting signal (aa 98127) to the in vivo targeting was examined directly using the GFP chimera aa 98127GFP chimera. With relatively low levels of expression, the aa 98127GFP construct (Fig. 7 A, yellow arrowhead) localized primarily to mitochondria visualized using an anti-COXI antibody (Fig. 7 B, yellow arrowhead). This is best visualized as yellow staining in the merged image (Fig. 7 C, yellow arrowhead). Like ROP2hc (Fig. 5, A, D, and G), increased accumulation of aa 98127GFP exhibited, in addition to colocalization with mitochondria (Fig. 7, DF, yellow arrowhead), evidence of clumping (Fig. 7 D) and localization to nonmitochondrial sites (Fig. 7, DF, green arrowhead). In addition, a subpopulation of mitochondria did not label with aa 98127GFP (Fig. 7, DF, red arrowhead).
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Characterization of ROP2hc binding to organelle membranes
Next, we compared the binding of the ROP2 derivatives tested in vivo, to purified organelles in vitro. Mitochondria and ER-enriched microsomes were prepared from murine livers (see Materials and methods). By immunoblot, the mitochondrial proteins COXI and COXIII were detected exclusively in the mitochondrial preparations (Fig. 8
A). Both anti-KDEL and anticalnexin antibodies strongly recognized the ER preparation (Fig. 8 A). Although no KDEL signal was visible in the mitochondrial preparation, a trace calnexin signal was apparent (Fig. 8 A), likely due to the mitochondrion-associated ER or mitochondrion-associated membrane (MAM) fraction, which is highly enriched in liver (Vance, 1990).
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All constructs, with the exception of GFP (Fig. 8 B, bottom panel), exhibited binding to mitochondria (Fig. 8 B, top three panels, carbonate). ROP2hc and 98127ROP2hcGFP were found almost exclusively in the mitochondrial pellet fraction (Fig. 8 B, top two panels, carbonate). Extraction with carbonate had no significant effect on ROP2hc binding, but did cause the displacement of some
98127ROP2hcGFP into the supernatant fraction (Fig. 8 B, top two panels, carbonate). These data suggest that despite the apparent absence of a predicted transmembrane domain, these proteins behave like integral membrane proteins in association with mitochondria. In contrast,
50% of aa 98127GFP bound to mitochondria (Fig. 8 B, third panel, carbonate -). Paradoxically, treatment with carbonate caused the fractionation of all of this construct into the pellet fraction (Fig. 8 B, third panel, carbonate +).
The binding of ROP2hc and 98127ROP2hcGFP to ER-enriched microsomes was tested. ROP2hc bound ER less efficiently than mitochondria, and while a portion was resistant to carbonate extraction, a significant pool remained in the supernatant fraction (Fig. 8 B, top panel, carbonate +). In contrast,
98127ROP2hcGFP, which targeted exclusively to ER (Fig. 6), exhibited efficient binding to ER in vitro that was completely resistant to extraction with carbonate (Fig. 8 B, second panel, carbonate +). The aa 98127GFP construct, containing the putative mitochondrial targeting domain of ROP2hc, exhibited very limited binding to ER (Fig. 8 B, third panel, carbonate -). As was observed with the binding of this protein to mitochondria, carbonate treatment caused aa 98127GFP to localize exclusively to the pellet fraction (Fig. 8 B, third panel, carbonate +). Given the highly basic nature of the aa 98127 sequence, precipitation in the presence of carbonate (pH 11.5) cannot be ruled out.
In light of the highly basic pI of ROP2hc (predicted pI 11.2) (Beckers et al., 1994), we were concerned that it too might pellet, albeit differentially, with mitochondria and ER compared with aa 98127GFP, after carbonate extraction as a result of its being precipitated. We tested this possibility using sucrose floatation gradients, in which precipitated ROP2hc should remain at the load fraction (Fig. 8 C, fractions 1 and 2), whereas membrane-associated ROP2hc should float to its equilibrium density. ROP2hc bound and floated to the appropriate equilibrium with both mitochondria (Fig. 8 C, Mito, NT) and ER (Fig. 8 C, ER, NT) cofractionating with the organelles as determined by Coomassie blue staining of the gels (unpublished data). Carbonate extraction failed to strip the protein from either mitochondria (Fig. 8 C, second panel, carbo) or ER (Fig. 8 C, bottom panel, carbo). These results indicate that full-length ROP2hc interacts with both mitochondria and ER with the characteristics of an integral membrane protein.
Given the high-affinity interaction of ROP2hc (Fig. 8, B and C) in vitro and the efficient trafficking of 98127ROP2hcGFP to ER in vivo, we examined whether these proteins could be imported cotranslationally into canine microsomes. Cotranslational import into the ER, based on protease protection, was not observed with either ROP2hc or
98127ROP2hc under conditions where Escherichia coli ß-lactamase (BLA) was efficiently imported, processed, and protected from exogenous protease (Fig. 8 D). Rather, combined with the results in Fig. 8, B and C, this suggests that ROP2hc and
98127ROP2hc form high-affinity interactions with the ER membrane.
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Discussion |
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Alternatively, ROP2hc could be using the low-efficiency bypass pathway (Pfaller et al., 1989) or a receptor analogous to the yeast TOM5 receptor. Both pathways are resistant to trypsin, with the TOM5 pathway capable of directing the translocation of the small Tim proteins (Tim 8, 9, 10, 12, and 13) into the IMS via the general insertion pore (Kurz et al., 1999). However, unlike the bypass pathway (Pfaller et al., 1989), import by the small Tim pathway occurs at 0°C and requires neither exogenous ATP nor the (Kurz et al., 1999).
Another class of proteins translocates across the plasma membrane into the cytoplasm of mammalian cells by a poorly understood mechanism involving the direct penetration through the lipid bilayer (reviewed in Schwarze and Dowdy, 2000). These proteins, containing protein translocation domains (PTD), include the HIV transactivator Tat (Frankel and Pabo, 1988), herpes simplex virus VP22 (Elliott and O'Hare, 1997), and the Drosophila melanogaster protein Antennapedia (Thoren et al., 2000). Although the mechanism is unclear (Schwarze and Dowdy, 2000), these molecules are highly basic, with a high concentration of arginine and/or twin arginine motifs implicated in diverse evolutionarily distant protein transduction pathways (Robinson, 2000; Schwarze and Dowdy, 2000). These features are all found in both ROP2hc, the NH2-terminal signal (aa 98127) (Fig. 3 A) required for mitochondrial targeting and import (Figs. 3 C and 7), and the remainder of the molecule (98127ROP2hc; aa 98465) that binds both mitochondrial and ER membranes (Fig. 8, B and C). Nonetheless, several important differences between the PTD-containing proteins and ROP2hc must be noted. First, unlike the PTD proteins that completely translocate across the bilayer (Schwarze and Dowdy, 2000), ROP2hc appears to insert into it without complete translocation (Figs. 3, 4, and 8). Secondly, while the translocation of PTD-containing proteins has not been examined in organelles, domains of ROP2hc exhibit distinct properties with regard to mitochondria and ER interaction (Fig. 8), suggesting that ROP2 may interact with specific protein or lipid domains in the target membranes.
The lack of complete translocation of ROP2hc, which does not encode obvious stop transfer sequences (von Heijne, 1990) into mitochondria in vitro, is best explained by the presence of a tightly folded domain in the protein. Folded protein domains block translocation across the MOM (Eilers and Schatz, 1986; Rassow et al., 1989, 1990). The presence of a folded domain in ROP2hc is inferred by the series of protease-resistant bands between 25 and 30 kD, after digestion of ROP2hc with either trypsin or proteinase K (Figs. 3 and 4). Notably, the presence of a translocation-incompetent subdomain within ROP2hc mimics the situation in vivo, where the ROP2 protein is embedded in the T. gondii PVM, making its complete translocation impossible.
The mechanism underlying the resistance of ROP2hc to carbonate extraction from either mitochondria or ER remains enigmatic. The protein lacks a stretch of noncharged and/or hydrophobic aa in the host cytoplasm exposed domain (aa 98465; Fig. 2 A) that are long enough to span a lipid bilayer (Beckers et al., 1994). The most likely mechanism involves direct insertion into the lipid bilayer, which may explain its interaction with both mitochondria and ER. Lipid and/or protein properties of these membranes also likely contribute to the differences in both the affinity of binding and the ability to translocate across the membrane (Figs. 3 and 8). ROP2hc inserts into the MOM (Figs. 3 and 4), integrating into it (Fig. 8, B and C). With ER membranes, high-affinity association (Fig. 8, B and C) is established without apparent translocation (Fig. 8 D). Whether the lack of microsomal import is merely a result of the in vitro conditions as has been suggested previously for TOM20 mutants (Kanaji et al., 2000) remains unclear.
ROP2 is the founding member of a family of highly related proteins found exclusively in T. gondii (Beckers et al., 1994, 1996; Sadak et al., 1988) that are likely to be essential. The likely essentiality and the presence of multiple isoforms (Beckers et al., 1994, 1996), complicates a gene knockout strategy to directly test the role ROP2 in PVMorganelle association. Furthermore, blockade of association after microinjection of anti-ROP2 antibodies is complicated by the existence of multiple targeting signals. Our previous data suggested that the extent of organelle association was governed by the concentration of organelles in the vicinity of the vacuole, and little else (Sinai et al., 1997).
The data establish a potential mechanistic paradigm by which a protein anchored in one organelle may "capture" another by inserting into its membrane, despite the apparent presence of a transmembrane domain. Such interactions, morphologically similar to PVMorganelle association, have been identified between mitochondria and other organelles, most notably the ER (reviewed in Bereiter-Hahn, 1990), and include the specialized MAM fraction of the ER (Trotter and Voelker, 1994; Vance and Shiao, 1996).
MAM-mediated ERmitochondrial association and the lipid trafficking, which is facilitated (Trotter and Voelker, 1994; Vance and Shiao, 1996), provides a potential paradigm for the role of PVMorganelle association in T. gondii biology. We have suggested that sites of PVM-associated organelles may be contact points for the bulk transfer of lipids in a manner analogous to MAM-mediated phospholipid transfer between mitochondria and ER (Sinai and Joiner, 1997; Sinai et al., 1997). Of note, preliminary experiments suggest that T. gondii is unable to synthesize phosphatidylcholine de novo (unpublished data), implying that parasite requirements for phosphatidylcholine and other lipids may be satisfied by scavenging from host cell sources.
In conclusion, the elucidation of a mechanism for PVMorganelle association highlights the importance of studying the cell biology of pathogenhost interactions. These studies reveal aspects of mammalian cell biology that would otherwise not be readily apparent. With millennia of coevolution behind them, intracellular pathogens have mastered cell biological principles that they are now revealing to us.
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Materials and methods |
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Cell culture and parasite infection
T. gondii strain RH, CHO-K1 (ATCC-CCL163), and primary human foreskin fibroblasts were maintained as described previously (Sinai et al., 1997). The kinetics of PVMmitochondrial association immediately after T. gondii infection were performed using human foreskin fibroblast cells preloaded with MitoTracker on coverslips in a 24-well dish placed in a 37°C water bath (Sinai et al., 1997). Cells were infected with T. gondii at a multiplicity of infection of 50. Parasite interaction with the cells was stopped at 1, 5, and 10 min postinfection by aspirating unbound organisms, washing once with PBS, and fixation in 3% paraformaldehyde. Coverslips were prepared for immunofluorescence using an anti-GRA3 antibody as described below.
Plasmid construction and PCR
Plasmid constructs used for in vitro expression were generated in pET17b (Novagen). The vectors used for in vivo expression of ROP2hc and GFP chimeras were pCR3.1(Invitrogen) and pWay2.1-Srf (Lo et al., 1998) (provided by Dr. Thom Hughes, Yale University College of Medicine, New Haven, CT), respectively. All inserts were generated using the PCR with either Taq (Perkin Elmer) or Vent polymerase (New England Biolabs, Inc.) according to the manufacturers recommendations, using pCBROP2.4 (15 ng) (Beckers et al., 1994) as a template. The sequences (5'3') of the forward (F) and reverse (R) PCR primers for ROP2hc cloned into pET17b were (F-ggaattccatattgagccacacagagactccgacaca, R-cgggtacctcagtggtggtggtggtggtgctggcggtagggggagc), and (F-aaaggtaccatgagccacacagagaccccgacacag, R-tttgaattcctacgttgggtggtgctggcggta) for pCR3.1 and pWay2.1-Srf. Inserts for ROP2hc80 and 98127ROP2hc subcloned into pET17b used the same reverse primer as ROP2hc, and the forward primers (aaaggtacc- cat0atgggaggctcatggctggag) and (ggaattccatatgtcagatggcggaggagaaccaccgcag), respectively. The inserts into pWay2.1-Srf generating the GFP chimeras aa 98127GFP and
98-127ROP2hcGFP were amplified using (F-aaaggtaccatgagccacacagagaccccgacacag, R-ataatacccgggtggtgcctggcggtaggggag) and (F-aaaggtaccatgtcagatggcgcaggagaaccaccg, R-tttgaattcctacgttgggtggtgctggcggta), respectively. Reverse primers used for ROP2hc and derivatives in pET17b contain a sequence encoding 4His codons after two His codons at aa 462 and 463 of ROP2 (Beckers et al., 1994). All primers were synthesized at the W.M. Keck Center (Howard Hughes Medical Institute, Yale University, New Haven, CT). Constructs cloned into pET17b, pCR3.1, and pWay2.1-Srf were ligated into NdeI-EcoRI, KpnI-EcoRI, and KpnI-SmaIrestricted gel-purified vectors, respectively, using sites engineered at the termini of the amplification primers. The presence of a T7 polymerase promoter in pCR3.1 and pWay2.1-Srf allows for in vitro protein synthesis. In vitro expression of BAP-ROP2TM/CT was achieved by subcloning that chimera from a pNTP/s derivative (Hoppe et al., 2000) into pCR3.1. Unless indicated, all enzymes for molecular biological application were purchased from New England Biolabs, Inc.
In vitro protein synthesis
35S-Metlabeled substrates for mitochondrial import assays were synthesized using a coupled rabbit reticulocyte lysate transcription/translation system (TNT; Promega), using 2 µg DNA per reaction. All constructs except that for human ornithine transcarbamylase (phOTC) (Horwich et al., 1984) were transcribed using the T7 polymerase (Promega). The OTC gene was transcribed using SP6 polymerase (Horwich et al., 1984) (Ambion or Promega).
Determination of ROP2 topology in the PVM
5 µl of in vitrosynthesized 35S-Metlabeled ROP2hc and BAP-ROP2TM/CT were immunoprecipitated in 200 µl IP using either 5 µl of anti-R/DG (see below) or 2 µl antibacterial alkaline phosphatase (anti-BAP) (3'5') along with 30 µl of 10% protein ASepharose (Pharmacia) for 4 h on a rotating wheel at 4°C. Protein ASepharose beads were harvested by centrifugation and washed thrice with IPW. The washed beads were boiled in SDS-PAGE sample buffer and eluted proteins resolved by SDS-PAGE, prepared for fluorography, and visualized as below.
Mitochondrial import assays
Mitochondria used in import assays were isolated from murine livers as described previously (Conboy and Rosenberg, 1981) and used immediately in import reactions. A standard protease protection assay was performed by incubating 8 µl of mitochondria (20 mg/ml in IB) with 12 µl of TNT reaction product for 20 min at 30°C. The reaction was split into three equal aliquots. The aliquots were treated with 0.6 µl of SBTI or PMSF (for trypsin or PK, respectively), 0.6 µl 10x protease (trypsin, 10 mg/ml or PK, 3.2 mg/ml) in IB, or 0.6 µl 10x protease + 0.6 µl 1% Triton X-100 in IB. Protease treatments used either trypsin (15 min at 4°C) or PK (30 min at 0°C). Reactions were stopped by adding either SBTI (10 mg/ml) or PMSF (1 mM final concentration) to all samples and incubating for 5 min at 4 or 0°C for trypsin- and PK-treated samples, respectively. Reactions were fractionated by centrifugation and prepared for SDS-PAGE as described in the figure legends. SDS-PAGE (Laemmli, 1970) was performed using either 13 (Fig. 3) or 15% (Fig. 4) Duracryl gels. Gels were prepared for fluorography using Autofluor (National Diagnostics) as recommended by the manufacturer, vacuum dried, and visualized using Kodak Biomax MR or AR x-ray film. To determine whether the NH2-terminal domain of ROP2hc and ROP2hc80 was protease protected, the mitochondrial pellet fraction from an import assay as above (both untreated and trypsin treated) was solubilized in 300 µl IP (IB + SBTI (10 mg/ml) and leupeptin (5 µM). The lysate was incubated with 5 µl of ascites fluid containing the mAb T34A7 and 15 µl of protein GSepharose (10% slurry in PBS) (Amersham Pharmacia Biotech). Immunoprecipitations were carried out essentially as described above.
Microsome import assays
Cotranslational import into canine microsomes was performed essentially as described by Hegde et al. (1998). E. coli BLA synthesized off an SP6 promoter was used as a positive control as described previously (Hegde et al., 1998). Both the canine microsomes and the pBLA construct were gifts from Dr. Ramanujan Hegde (National Institutes of Health, Bethesda, MD).
Binding and fractionation assays
Murine liver mitochondria (Conboy and Rosenberg, 1981) and ER (Paulik et al., 1988) were prepared as described previously and frozen at -80°C until use. The purity of the preparations was assessed using immunoblot analysis with organelle-specific markers (COXI and COXIII) for mitochondria (1:1,000]), calnexin (1:500), and anti-KDEL (1:1,000 for ER). Immunoblots were performed as previously described (Sinai et al., 1997).
Labeled ROP2hc, 98127ROP2hcGFP, aa 98127GFP, and GFP were synthesized in vitro, added in duplicate tubes to either purified mitochondria or ER (100 µg total protein), and brought to 100 µl with Tris-buffered isosmotic sucrose, pH 7.4. Binding was allowed to proceed in ice water for 30 min. Carbonate treatment was performed by the addition of 100 µl of 0.2 M sodium carbonate, pH 11.5 for 30 min. The untreated controls received an equal volume of isosmotic sucrose. After incubation, the samples were centrifuged for 15 min in an air-driven ultracentrifuge (Airfuge; Beckman Coulter) set at 30 psi. Both the organelle pellets and the supernatants were recovered and resolved by SDS-PAGE.
To confirm the membrane association of ROP2hc, sucrose floatation gradients were performed. Samples prepared as above were incorporated into 1.5 ml of 60% sucrose in 10 mM Tris-Cl, pH 7.4, and gradients established as described previously (Sinai et al., 1997). In light of dilution with the sample, the sucrose concentration in the load fraction was 55%. The gradients were centrifuged and harvested as described previously (Sinai et al., 1997). 150 µl from each fraction was mixed with 4x SDS-PAGE sample buffer, and boiled and resolved on a 10% Duracryl gel. The gels were stained by Coomassie blue to identify the organelle-positive fractions, treated with a fluorophore (Amplify; National Diagnostics), and exposed for fluorography as above to identify ROP2hc-positive fractions.
Antibodies
Antibodies used in this study for immunofluorescence microscopy and/or immunoblot analysis include mAbs T34A7 and T3H11 against ROP2,3,4 (Sadak et al., 1988) and GRA3 (Bermudes et al., 1994), respectively, anti-mammalian COXI (Molecular Probes), anti-cytochrome c (Zymed Laboratories), and anti-KDEL (StressGen Biotechnologies). In addition, rabbit sera against GFP (CLONTECH Laboratories, Inc.), mammalian COXIII (provided by Dr. Pietro De Camilli, Yale University School of Medicine) (Nemoto and De Camilli, 1999), calnexin (Hebert et al., 1995), E. coli alkaline phosphatase (BAP) (3'5'), and the R/DG fraction of T. gondii (Beckers et al., 1994) were used. Finally, a chicken anti-TOM20 IgY was provided by Drs. Ing Swie Goping and Gordon Sinore (McGill University, Montreal, Canada) (Goping et al., 1995).
Transfection
CHO cells, at 80% confluence in a six-well plate (Falcon), were transfected using the QIAGEN Superfect reagent as recommended by the manufacturer. Cells were incubated with the transfection mix for 3 h at 37°C, washed once with PBS, trypsinized, and plated onto eight coverslips, each placed in a 24-well culture dish. The transfected gene product was detected by immunofluorescence analysis between 18 and 24 h posttransfection.
Fluorescence Microscopy
Transiently transfected cells were seeded on coverslips and immunofluorescence was performed as described previously (Sinai et al., 2000). All antibodies were diluted in 20% goat serum as follows: mouse monoclonals T34A7 and T3H11 at 1:300, anti-COXI, anti-cytochrome c at 1:600, anti-KDEL at 1:200, and rabbit anti-GFP, anti-COXIII, and anti-calnexin at 1:500. Immunofluorescence on MitoTracker-labeled cells (described in Sinai et al., 1997) was performed using either T34A7 or anti-GFP incubation for at least 1 h at room temperature. Double labeling (T34A7 with anti-COXIII or anti-calnexin, anti-GFP with anti-COXI or anti-cytochrome c, or anti-KDEL) was performed simultaneously for at least 1 h at room temperature. Species-specific secondary antibodies conjugated to either FITC (Calbiochem), Oregon green (Molecular Probes) or Texas red (Molecular Probes) were used as indicated in the figure legends.
Laser scanning confocal microscopy was performed at the Electron Microscopy and Imaging Suite at the University of Kentucky College of Medicine using a Leica TCS True Confocal microscope system. All images were acquired using a 100X/1.4 NA Plan Apo oil immersion objective. Digitized images were imported into Adobe Photoshop, and all adjustments for brightness and contrast applied uniformly to the entire field.
Electron Microscopy
Electron microscopy on CHO cells infected with T. gondii for 20 h was performed as described previously (Sinai et al., 1997).
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Footnotes |
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Acknowledgments |
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This work was supported by a postdoctoral affiliate fellowship (No. 98-2002T) from the American Heart Association, New Faculty Startup funds from the University of Kentucky Research Challenge Trust Fund (RCTF), an Institutional Research grant (IRG 85-001-13-IRG) from the American Cancer Society to A.P. Sinai, and a National Institutes of Health grant (AI30060) and a Burroughs Wellcome Fund Scholar Award in Molecular Parasitology to K.A. Joiner.
Submitted: 22 January 2001
Revised: 17 May 2001
Accepted: 1 June 2001
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