Department of Plant Sciences, The George S. Wise Faculty of Life Sciences, Tel Aviv University, Tel Aviv 69978, Israel
* Author for correspondence (e-mail: aviah{at}post.tau.ac.il)
Accepted 14 April 2005
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Summary |
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Key words: Cell cycle, Cry1C, Bacillus thuringiensis, Lipid rafts, Caveolin, Resistance
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Introduction |
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Glycosylphosphatidylinositol (GPI)-anchored amino-peptidase N (APN) and cadherin-like proteins (CLPs) were identified as receptors of Cry proteins in various lepidopteran larvae. Although interaction of APNs with different Cry1A proteins were clearly demonstrated in vitro (Banks et al., 2001; Garner et al., 1999
; Gill et al., 1995
; Knight et al., 1994
; Luo et al., 1999
; Nakanishi et al., 1999
; Oltean et al., 1999
; Rajagopal et al., 2003
; Simpson and Newcomb, 2000
; Yaoi et al., 1997
), APN capacity to mediate toxicity in vivo has only been shown by applying double-strand RNA to Spodoptera litura larvae or by ectopic expression in Drosophila melanogaster larvae (Gill and Ellar, 2002
; Rajagopal et al., 2002
). Other attempts only showed binding of the membrane-assembled foreign APN to the Cry toxin, without causing toxicity (Banks et al., 2003
; Denolf et al., 1997
; Luo et al., 1999
; Rajagopal et al., 2003
; Simpson and Newcomb, 2000
). By contrast, Cry1Aa sensitivity was achieved by expressing Bombyx mori CLP in human HEK293 cells (Tsuda et al., 2003
) and also in Sf9 cells (Nagamatsu et al., 1999
) but not in murine COS7 cells where it was ineffective (Tsuda et al., 2003
). Moreover, natural transposon insertion in the gene encoding CLP led to Cry1Ac resistance (Gahan et al., 2001
; Morin et al., 2003
).
These observations indicated that APN and CLPs are not the only components involved in mediating Cry protein toxicity. Consequently, the presence and correct assembly of other membrane components has recently been proposed as a prerequisite for Cry1A toxicity. APN localization to lipid raft domains of the plasma membrane in epithelial cells of Heliothis virescens and Manduca sexta larval midgut lends support to this approach (Zhuang et al., 2002). Lipid rafts are membrane micro-domains enriched in GPI-anchored proteins, sphingolipids and sterols, and are defined by their insolubility in Triton X-100 at low temperature (Brown and Rose, 1992
; Zurzolo et al., 2003
). Clustering and recruitment of other proteins in lipid rafts represent dynamic processes that occur in the membrane in response to various signals upon interaction with ligands (Manes et al., 2003
). Although lipid raft integrity was shown to be essential for efficient binding of Cry1A (Zhuang et al., 2002
), the sequence of events that includes interaction with raft-associated APN and non-raft-associated CDRs awaits further elucidation.
Spodoptera-frugiperda-derived Sf9 cells are highly and specifically sensitive to Cry1C toxin (Kwa et al., 1998; Rang et al., 1999
; Vachon et al., 1995
; Villalon et al., 1998
). Cry1C pore-formation in Sf9 cells was verified by evaluating changes in membrane permeability and cellular shape (Guihard et al., 2001
; Guihard et al., 2000
; Vachon et al., 1995
; Villalon et al., 1998
). Sf9-specific response to Cry1C, was utilized to verify specificity and toxicity of Cry1C-Cry1A chimeric fusions (Rang et al., 1999
) and mutated Cry1C proteins (Smith and Ellar, 1994
; Tayabali and Seligy, 1995
). Although Sf9 cells have been intensively studied as an insect model system, membrane components involved in the interaction with Cry1C had not been characterized.
The present study elucidates cell-cycle-dependent Cry1C-insensitivity in Sf9 cells, showing Cry1C tolerance during mitosis that gradually disappears in G1 phase. Arresting Sf9 cells in G2-M phase caused the same loss of Cry1C sensitivity. The reduced Cry1C-binding-capacity and -sensitivity during G2-M phase was correlated with the inability to isolate defined lipid rafts from G2-M cells, suggesting that a correct membrane lipid raft organization, accomplished during G1 phase and abolished in M phase, is essential for Cry1C interaction.
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Materials and Methods |
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Sf9 propagation and photography
The Spodoptera frugiperda Sf9 cell line (ATCC CRL-1711) was generated from the IPLB-Sf21 cell line, which originated from pupal ovarian tissue (Vaughn et al., 1977). Sf9 cells were grown in Grace's medium (Biological Industries, Beit Haemek, Israel) supplemented with 10% (v/v) heat-inactivated fetal calf serum at 27°C. The cells were diluted in 12-well plates and Cry1C (500 ng/ml) was added 24 hours later during log-phase stage. Cell swelling and cell death were detected with an Olympus BX52 microscope under visible or fluorescent light by staining with 0.2 µM ethidium homodimer-1 dye (Fluka, excitation
495 nm, emission
635 nm). Cells were photographed every 3 minutes by using the Till Photonics fluorescence imaging system and the TillvisION computer program.
Cell synchronization
Cell cultures were synchronized using nocodazole (Sigma), a G2-M phase arresting agent. Nocodazole (10 µg/ml) (Braunagel et al., 1998) was added to the cells 24 hours after culture dilution. The response to Cry1C was evaluated 24 to 48 hours after nocodazole addition. Nocodazole was removed by three consecutive washes with growth medium.
Assaying Cry1C sensitivity
Normal or G2-M-arrested adherent Sf9 cells were grown in Grace's medium with serum. When a density of 1000 cells per mm2 surface was reached, increasing concentrations (100 to 5000 ng/ml) of Cry1C were added to the medium for 4 hours and cells were incubated at 27°C. Dead cells were distinguished from toxin-unaffected living cells by their pre-rupture swollen shape or their post-rupture shrunken-shape. Dead cells also had granular cytoplasm, allowing definite identification, counting and LC50 estimation. Staining with ethidium homodimer-1 was an additional way to identify dead cells when recovery from the nocodazole treatment was monitored by fluorescence-microscopy.
Membrane lipid rafts purification
Lipid rafts isolation (Lavie et al., 1998) was performed in a cold room (4°C). Sf9 and G2-M arrested cells were grown in 72-cm2 flasks to a density of 30 x106 each. Cells were washed twice with ice-cold PBS (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4). Ice-cold lysis buffer (1 ml) [25 mM 2-N-morpholinoethane-sulfonic acid (MES, Sigma) pH 6.5, 150 mM NaCl, 1% Triton X-100, 1 mM sodium pyrophosphate, 1 mM sodium vanadate, 1 mM PMSF, 10 µg/ml aprotynin, 10 µg/ml leupeptin] was added to each flask and the lysed cells were scraped from the surface. The lysate was collected and the final volume was adjusted to 1.5 ml with ice-cold lysis buffer. One milliliter of homogenized (15 strokes in Dounce homogenizer) lysate was mixed with an equal volume of 80% sucrose in MES buffer (25 mM MES pH 6.5, 150 mM NaCl) in SW41 tubes. Each tube was stepwise overlaid with 4 ml of 30% sucrose and 4 ml of 5% sucrose dissolved in MES buffer. The samples were centrifuged at 4°C for 20 hours in an SW41 rotor (Beckman) at 175,000 g. Fractions of 900 µl were collected from the top. Lipid rafts appeared in the middle of the gradient (fractions four to six) and soluble membrane proteins in fraction 11. Lipid raft and soluble fractions (30 µl/fraction) were analyzed by SDS-PAGE and western blotting. Before loading on gels, lipid raft fractions of G2-M arrested cells were concentrated 30-fold by acetone precipitation.
Antibody (Ab) against human caveolin-1 (Santa Cruz Biotechnology, N-20 sc-894 affinity purified rabbit polyclonal Ab raised against the N terminal region of human caveolin-1) was used to identify Sf9 caveolin that served as a marker for lipid raft proteins.
Western analysis of total proteins
Caveolin was identified in western blots by comparing responses of the anti-human caveolin 1 polyclonal Abs with total proteins extracted from Sf9 cells and caveolin-overexpressing human HT29MDR cells (Lavie et al., 1998). Total proteins were prepared by resuspending PBS-washed cell pellet in 500 µl extraction buffer [50 mM Tris-HCl, pH 7.8, 2% SDS, 1 mM phenyl methyl sulfonyl fluoride (PMSF), 10 mg/ml aprotynin, 10 mg/ml leupeptin]. After a 15-minute incubation at 4°C and two cycles of 10-second sonication (30% in Sonic-Vibra Cell) followed by centrifugation (14,000 g), 0.16 volumes of basic sample buffer (62.5 mM Tris, pH 11, 10% SDS, 50% glycerol, 5% ß-mercapthoethanol, 0.06% Bromophenol-Blue) was added to the supernatant. The samples were then heated in boiling water for 10 minutes. Total proteins (50 µg per lane) were separated on SDS-PAGE (15% acrylamide) and, after blotting onto Immobilon membranes (Amersham Bioscience), were incubated with rabbit anti-human caveolin 1 Abs (sc 894) for 12 hours and followed by horseradish peroxidase-conjugated goat anti-rabbit IgG (Jackson, 115-035-003) for 1-2 hours. The signal was detected by enhanced chemiluminescence (ECL) (Amersham Biosciences). Caveolin-specific recognition by anti-caveolin 1 Abs was shown by pre-incubating the Abs with a blot containing equal amounts (100 µg) of HT29MDR total proteins.
Cry1C binding
Normal or G2-M-arrested Sf9 cells (5 x105 cells) were mixed with 0.5 ml serum-supplemented Grace's medium containing 1 µg/ml or 2.5 µg/ml Cry1C. Binding reactions were incubated in the dark to avoid nocodazole degradation. After 5 or 90 minutes, reactions were stopped by centrifugation and the cells were washed twice with PBS. Washed cells were suspended in sample buffer, boiled and then equal amounts of total protein (9 µg/lane) were separated on SDS-PAGE (10%). Bound Cry1C was identified by western blotting using anti-Cry1C polyclonal Abs. The NIH Image program (US National Institutes of Health, http://rsb.info.nih.gov/nih-image/) was used to quantify scanned immunoblot bands. The same blots were re-probed with a mouse anti-chicken actin Ab (ICN, 69100) followed by alkaline phosphatase-conjugated goat anti-mouse IgG (Sigma, A3562), to measure total protein content in the loaded samples.
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Results |
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G2-M-phase arrested Sf9 cells are Cry1C-tolerant
Analysis of Cry1C-treated cells indicated that dividing Sf9 cells were insensitive to Cry1C at M phase and early G1 phase (Fig. 1). To obtain G2-M synchronized cells, nocodazole (10 µg/ml), an inhibitor of tubulin polymerization, was added to the culture. Cry1C-sensitivity tests were performed by exposing normal (untreated) and G2-M-arrested Sf9 cells to various concentrations (from 0.1 to 5.0 µg/ml) of Cry1C (Fig. 2). Exposure to Cry1C for 4 hours killed the non-arrested cells and reached complete mortality at 250 ng/ml, whereas the G2-M cells started to be affected at higher Cry1C concentrations, above 500 ng/ml. Consequently the recorded LC50 of G2-M cells was 5-fold higher than that of the normal cells (Fig. 2B). This reduction in Cry1C-sensitivity can be attributed to the presence of fewer membrane receptors or/and lower membrane-binding-capacity during mitosis.
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Cry1C insensitivity is typical to M phase and disappears during G1 phase
The ability of G2-M arrested cells to regain Cry1C-sensitivity was examined after the removal of nocodazole by three consecutive changes of the growth medium. A gradual release of the arrested cells from the `nocodazole barrier' occurred, allowing cells to complete mitosis, recover and grow similarly to untreated cells. The recovery of nocodazole-treated cells lasted several hours (Braunagel et al., 1998) with the appearance of the first cell divisions only 6 hours after washing off the nocodazole (data not shown).
To investigate the regained Cry1C-sensitivity, G2-M cells were exposed to Cry1C (300 ng/ml) immediately after the removal of nocodazole (Fig. 3A,C). In parallel treatments, normal cells (Fig. 3A, panels labelled `Unarrested') and G2-M cells (Fig. 3B) were both exposed to Cry1C and served as controls for Cry1C activity. Cellular nucleic acids in dead cells stained with ethidium homodimer-1 served as an indicator of cell mortality. In normal cells the interaction of the dye started immediately after Cry1C application reaching 100% staining, which reflected mortality within 5-6 hours (Fig. 3A,C). By contrast, cells that were released from the G2-M arrest remained viable during this 6-hour recovery period from the nocodazole effect, and regained Cry1C-sensitivity only after about 9 hours (Fig. 3A,C). Recovered cells became gradually sensitive to Cry1C and died evident by the appearance of more red-stained cells after 12-18 hours of exposure to Cry1C (Fig. 3A,C and supplementary material Movie1). Cells that remained at continuous G2-M arrest stayed alive throughout the Cry1C treatment and showed a mortality rate of only 3% (Fig. 3B,C). These observations clearly demonstrated that Sf9 cells become Cry1C-tolerant only during G2-M transition.
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Whereas lipid rafts could be easily isolated from normal cells (Fig. 4 lanes 2-4), almost no lipid raft proteins were identified in the G2-M cells (Fig. 4 lanes 5-7) by either silver-staining or by probing with the anti-Cav-1 Ab; not even in the 30-fold concentrated lipid raft fractions. By contrast, defined caveolin bands were identified in the Triton X-100 soluble membrane fractions of the G2-M cells (Fig. 4, lane 8), indicating that caveolin was still present in the plasma membrane but shifted to the membrane-liquid-disordered phase (Simon and Ehehait, 2002). The absence of Triton X-100-insoluble lipid rafts in G2-M-arrested Sf9 cells coincided with the appearance of Cry1C-tolerance. Therefore, these results strongly suggest that the disappearance of membrane lipid raft structures from the plasma membrane during mitosis is the cause for the transient loss of Cry1C sensitivity.
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Discussion |
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In this study, analysis of Cry1C toxicity to Sf9 cells was under optimal growing conditions (Grace's medium supplemented with serum) to avoid any stress-induced physiological changes (Doverskog et al., 2000; Drews et al., 1995
). In other studies, Sf9-cell susceptibility to pore formation by Cry1C varied according to the molecular and ionic content of the medium (Guihard et al., 2000
; Villalon et al., 1998
). Sf9 cells that were kept in isotonic buffers were affected by lower concentrations of Cry1C (Kwa et al., 1998
), compared with cells grown in supplemented Grace's medium.
Within 4 hours, Sf9 cells that were grown at these optimal growth conditions and then treated with 300 ng/ml Cry1C, showed a mortality rate of 98-99%. However, neonate S. frugiperada larvae that possess relatively high midgut-proteolysis-capacity, only showed a mortality rate of 20% after treatment with a similar concentration of Cry1C (M. Adang, University of Georgia, Athens, GA, personal communication). Membrane changes that occurred during mitosis might prevent Cry1C-toxin-binding and Cry1C toxicity. Relatively high Cry1C concentrations, in the range of 1000-5000 ng, were required to affect G2-M cells (Fig. 2), probably owing to reduced binding capacity caused by M-phase-linked membrane rearrangements. Such rearrangements were evident by the correlative lack of lipid rafts observed in G2-M cells (Fig. 4) and the appearance of lipid raft components such as caveolin in the soluble membrane fractions. Membrane-changes accompanied by lipid raft rearrangements were also observed during mitosis in yeast cells (Bagnat and Simons, 2002; Rajagopal et al., 2003
; Wachtler et al., 2003
). In dividing yeast cells, lipid rafts as detected by filipin fluorescence were confined to the vicinity of the contractile ring; during the interphase stage, however, lipid rafts were spread over the yeast cell tips. Another rearrangement of lipid rafts in yeast is induced during mating (Bagnat and Simons, 2002
), resulting in clustering of lipid rafts that contain mating-related proteins in the mating projection. A mutant with reduced lipid raft levels showed impaired mating, suggesting a direct correlation between the assembly of lipid rafts and mating. No membrane rafts were isolated from dividing Sf9 cells, which indicates a specific M-phase rearrangement of lipid raft components; this was evident by the appearance of caveolin-1-like protein in the membrane-soluble fraction.
Lipid rafts are considered to be highly dynamic entities that continually change their size and composition, and are also able to coalesce to larger clustered structures (Lucero and Robbins, 2004). Thus, the M-phase-dependent lack of defined lipid raft domains, which correlated with Cry1C-insensitivity, might reflect transient lipid rafts de-clustering during cell division.
Hence, these data extend the currently existing evidence about the involvement of lipid rafts in Cry toxicity. The well-defined Cry1A receptor APN was shown to be a lipid-raft-resident protein anchored to the raft structure by its GPI moiety (Zhuang et al., 2002). Recently, an APN homologue has been identified in the midgut epithelium of S. litura larvae as a potential Cry1C receptor based on its interaction with the toxin in vitro and on RNAi silencing experiments carried out in living S. litura larvae (Agrawal et al., 2002
; Rajagopal et al., 2002
). These data indirectly suggest the involvement of lipid rafts also in Cry1Cmembrane interaction. Thus it is proposed that, during M phase, the receptors and probably other, as yet unidentified, components essential for Cry1C intoxication are not recruited to the lipid rafts and, consequently, the interaction of the toxin with the membrane is interrupted. Lipid raft clustering is also an essential step in aerolysin pore-formation and in the anthrax tripartite-toxin entry. An initial interaction with the cell membrane induces further clustering of lipid rafts because it is an essential step in both processes (Abrami et al., 2003a
; Abrami et al., 2003b
).
Understanding the mechanisms of resistance to Cry proteins is very important for the future use of Bt toxin in agriculture. As yet, no significant development of resistance has been reported in Bt crops (Carriere et al., 2003; Morin et al., 2003
), although field resistance of Plutella xylostella was generated by Bt sprays (Shelton et al., 1993
). Cry-resistant populations have been selected by exposing various lepidopteran larvae to high concentrations of specific Cry proteins in the laboratory (Ferre and Van Rie, 2002
). Naturally mutated host genes encoding a cadherin-like receptor (Gahan et al., 2001
; Morin et al., 2003
) or artificially mutated genes encoding ß-1,3-glycosyltransferase (Griffitts et al., 2001
) were identified as causal genes leading to Bt-resistance. Additional resistance mechanisms described so far were attributed to the loss of larval midgut proteases that are required to activate protoxins (Ferre and Van Rie, 2002
; Oppert et al., 1997
) or to higher gut-proteolytic-activity that might lead to toxin degradation (Loseva et al., 2002
). Recently, reduction in binding-affinity that depends on membrane integrity has also been proposed to play an important role in larval resistance to Cry proteins (Avisar et al., 2004
; Ferre and Van Rie, 2002
).
This study elucidates the dependence of Cry1C-binding and Cry1C-toxicity on membrane lipid rafts, and shows that these lipid raft domains are absent during cell division. Cell-cycle-dependent Cry1C-resistance, as demonstrated in Sf9 cells, represents a completely different perspective of resistance to Cry proteins and emphasizes the crucial role of membrane organization. In the gut epithelium of lepidopteran larvae, stem cells exist that maintain a high cell-division-frequency and give rise to mature Goblet and columnar cells (Loeb et al., 2003; Loeb et al., 2001
). Upon exposure to Cry toxin, these dividing cells might be less susceptible than differentiated epithelial cells and might serve as a reservoir for producing undamaged mature cells during recovery from treatments with Cry toxin. Further analysis of membrane changes during the M phase, and their relation to, as yet undefined, Cry1C-interacting membrane components will clarify the role of the cell cycle in the development of Cry-protein resistances.
The observed lipid raft rearrangement that occurs during the G2-M phase in Sf9 cells may be common to many other dividing cells and, therefore, may affect many lipid-raft-involving interactions. Lipid rafts have been implicated in regulating numerous cellular processes including exocytosis (Salaun et al., 2004), intracellular trafficking of proteins, toxins and pathogens that require specific interactions with lipid raft resident proteins and sphingolipids (Helms and Zurzolo, 2004
; Taieb et al., 2004
), and many signaling pathways (Zhai et al., 2004
). Mobile changes in raft organization, including shifts of small rafts into clustered platforms, are essential for normal cellular processes such as polar apical sorting and many other processes that depend on the recruitment to the rafts of proteins with weak raft affinities (Fullekrug and Simons, 2004
). Therefore, the lack of lipid raft domains in dividing cells might affect many endogenous cellular processes as well as communication with the external environment. Thus within a limited time window, dividing cells may exhibit different communication abilities compared to differentiated cells at the interphase stage.
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Acknowledgments |
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Footnotes |
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References |
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