Cell-cycle-dependent resistance to Bacillus thuringiensis Cry1C toxin in Sf9 cells

Dror Avisar, Michal Segal, Baruch Sneh and Aviah Zilberstein*

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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The Sf9 cell line, derived from the moth Spodoptera frugiperda, is highly and specifically sensitive to the Bacillus thuringiensis Cry1C toxin. Upon exposure to Cry1C, ionic pores are formed in the plasma membrane leading to cell swelling and death. Here, we describe a unique transient tolerance to Cry1C of dividing cells, which allowed completion of the division process in the presence of Cry1C. Correlatively, arresting the cells at G2-M phase by nocodazole treatment rendered them insensitive to Cry1C. When the arresting agent was removed, the cells completed their division and gradually regained Cry1C sensitivity. In comparison to normal cells with 1-2% cell-division frequency, the M-phase arrested cells bound less toxin in binding assays. Moreover, no lipid rafts could be isolated from the membranes of M-phase arrested cells. Caveolin-1, identified here for the first time in insect cells, was immunodetected as a lipid raft component of normal cells, but was only present in the membrane-soluble fraction of G2-M-arrested cells. Thus M-phase-linked changes in lipid raft organization may account for diminished Cry1C binding and toxicity. Furthermore, considering the pivotal role of lipid rafts in different cell functions of many cell types, the lack of organized lipid rafts in dividing cells may transiently affect cell susceptibility to pathogens, toxins and other lipid raft-linked functions.

Key words: Cell cycle, Cry1C, Bacillus thuringiensis, Lipid rafts, Caveolin, Resistance


    Introduction
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 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Bacillus thuringiensis (Bt) crystal proteins (Cry) are natural insecticides that are intensively used in agriculture as proteins being produced in formulated bacteria or constitutively synthesized in transgenic crops. The Cry protoxin is dissolved and cleaved by the insect's trypsin-like gut proteases to form the N-terminal active toxin that interacts with specific cell membrane receptors. Non-selective ion channels are then formed, leading to cell death by disrupting membrane-selective permeability and cellular electrolyte-balance (Schnepf et al., 1998Go).

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., 2001Go; Garner et al., 1999Go; Gill et al., 1995Go; Knight et al., 1994Go; Luo et al., 1999Go; Nakanishi et al., 1999Go; Oltean et al., 1999Go; Rajagopal et al., 2003Go; Simpson and Newcomb, 2000Go; Yaoi et al., 1997Go), 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, 2002Go; Rajagopal et al., 2002Go). Other attempts only showed binding of the membrane-assembled foreign APN to the Cry toxin, without causing toxicity (Banks et al., 2003Go; Denolf et al., 1997Go; Luo et al., 1999Go; Rajagopal et al., 2003Go; Simpson and Newcomb, 2000Go). By contrast, Cry1Aa sensitivity was achieved by expressing Bombyx mori CLP in human HEK293 cells (Tsuda et al., 2003Go) and also in Sf9 cells (Nagamatsu et al., 1999Go) but not in murine COS7 cells where it was ineffective (Tsuda et al., 2003Go). Moreover, natural transposon insertion in the gene encoding CLP led to Cry1Ac resistance (Gahan et al., 2001Go; Morin et al., 2003Go).

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., 2002Go). 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, 1992Go; Zurzolo et al., 2003Go). 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., 2003Go). Although lipid raft integrity was shown to be essential for efficient binding of Cry1A (Zhuang et al., 2002Go), 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., 1998Go; Rang et al., 1999Go; Vachon et al., 1995Go; Villalon et al., 1998Go). Cry1C pore-formation in Sf9 cells was verified by evaluating changes in membrane permeability and cellular shape (Guihard et al., 2001Go; Guihard et al., 2000Go; Vachon et al., 1995Go; Villalon et al., 1998Go). Sf9-specific response to Cry1C, was utilized to verify specificity and toxicity of Cry1C-Cry1A chimeric fusions (Rang et al., 1999Go) and mutated Cry1C proteins (Smith and Ellar, 1994Go; Tayabali and Seligy, 1995Go). 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.


    Materials and Methods
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 Materials and Methods
 Results
 Discussion
 References
 
Purification of Cry1C toxin
Cry1C toxin from B. thuringiensis subsp. aizawai was produced and purified as previously described (Guihard et al., 2000Go). Protein concentration was determined using bovine serum albumin as a standard.

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., 1977Go). 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., 1998Go) 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., 1998Go) 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., 1998Go). 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.


    Results
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 Materials and Methods
 Results
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 References
 
Sf9 cells are relatively Cry1C-insensitive during mitosis and early G1 phase
Cry1C induces the formation of non-selective ion channels in the cell membrane (Schwartz et al., 1991Go; Vachon et al., 1995Go) leading to cell-swelling and finally the collapse of the cell. To verify the time course of the toxic effect, Cry1C (500 ng/ml) was applied to Sf9 cells during continuous light-microscopy screening of adherent cells in log-phase growth. Such Cry1C concentration killed about 98-99% of the cells. Most of the cells were gradually killed by the toxin in the first 4 hours, except for dividing cells – making up 1.58±0.95 percent of the cell population – that were not affected (Fig. 1). The two dividing cells shown in Fig. 1A completed division within 36 minutes and the resulting daughter cells remained viable even after 450 minutes while all the neighboring non-dividing cells had already been killed by the toxin. Fig. 1B demonstrates a single cell that entered M phase relatively late, after more than 210 minutes of incubation with Cry1C, yet could complete division within 456 minutes exposure to the toxin; its daughter cells remained alive even after 22 hours. The daughter cells that could survive during early G1 phase became gradually Cry1C-sensitive with cell-cycle progression, whereas those resulting from late occurring divisions (as shown in Fig. 1B) remained alive rather longer.



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Fig. 1. Sf9 cells gain temporary Cry1C-tolerance during mitosis and early G1 phase. Log-phase Sf9 cells were incubated with 500 ng/ml Cry1C and photographed every 3 minutes. (A,B) Two microscopic fields with dividing cells (indicated by arrows) that were Cry1C-insensitive and progressed in their divisions. Inserts show the magnified regions (x2.3) to their right. After division and in the presence of Cry1C, daughter cells remained alive at least 300 minutes (A) or up to 22 hours (B). The other non-dividing cells gradually died within 180 minutes. A mean value of dividing cells in percent (1.58±0.95) was estimated by counting 10 different cell cultures.

 

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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Fig. 2. G2-M-arrested Sf9 cells survived treatment with Cry1C. Normal Sf9 cells and G2-M-arrested Sf9 cells were exposed to increasing concentrations of Cry1C for 4 hours. Dead and viable cells were counted (three repeated experiments). Dose-dependent mortality curves (A) were used to estimate LC50 (B). The control curve, data from untreated cells exposed to a mixture of Cry1C (increasing concentrations) and nocodazole (10 µg/ml), indicates that nocodazole has no effect on Cry1C-activity.

 

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., 1998Go) 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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Fig. 3. Sf9 cells gradually regain Cry1C-sensitivity while progressing through the cell-cycle and recovering from G2-M arrest. (A) Normal (unarrested) Sf9 cells (left) and Sf9 cells released from the G2/M arrest after nocodazole had been washed off (right) and (B) G2-M-arrested cells. All cells were exposed to 300 ng/ml Cry1C for 18 hours. Dying cells with a disrupted plasma membrane were detected by staining nucleic acids with ethidium homodimer-1 (red fluorescence). (C) Time course of Cry1C-induced cell death in the three treatments shown in A and B. Most of the untreated Sf9 cells (90%) died within 6 hours of Cry1C-exposure ({bullet}). Together with cell-cycle progression in cells released from G2-M arrest, a gradual sensitivity to Cry1C was regained, detected by dead red-stained cells that started to appear after 8-9 hours of Cry1C exposure ({square}). Very low cell mortality was observed in G2/M-arrested cells following application of Cry1C ({blacktriangleup}).

 
Lipid rafts in normal cells and in G2-M-phase arrested Sf9 cells
The involvement of lipid rafts in Cry1A toxicity was recently reported (Zhuang et al., 2002Go). To examine the link between the presence of membrane lipid rafts and Cry1C-sensitivity, lipid rafts were isolated from normal-growing and G2-M-arrested Sf9 cells. Five different lipid raft preparations were analyzed by SDS-PAGE followed by either silver staining (Fig. 4B) or immunoblot analysis with anti-human caveolin-1 (Cav-1) Abs (Fig. 4A). The caveolin-1 oligomer is a well defined lipid raft component of mammalian caveolae (Anderson, 1998Go; Liu et al., 2002Go; Simons and Toomre, 2000Go). Caveolin orthologues were also described in the invertebrate Caenorhabditis elegans (Tang et al., 1997Go). In the lipid raft fractions of the normal Sf9 cells and human HT-29-MDR cells (Fig. 4A, lanes 2-4 and lane 1, respectively), the anti human Cav-1 Abs recognized bands corresponding to putative caveolin oligomers. To further validate Sf9 caveolin recognition by the Abs, the immunodetected pattern of total proteins extracted from Sf9 cells and human HT-29-MDR cells that overexpress caveolin-1 (Lavie et al., 1998Go) were compared (Fig. 4C). According to the gel-migration pattern, mammalian caveolins are shorter than Sf9 caveolin monomers. The latter is similar to C. elegans caveolin 1, whose predicated molecular mass is 26,293 Da (Tang et al., 1997Go). Such recognition of the insect orthologues, described here for the first time, indicated the existence of conserved epitopes of caveolins in distantly related organisms.



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Fig. 4. Comparison of lipid rafts of untreated and G2-M-arrested Sf9 cells. Lipid rafts were isolated from unarrested and G2-M arrested Sf9 cells (lanes 2-4 and 5-7, respectively). The lipid raft fractions of G2-M cells were concentrated 30-fold after isolation. (A) Immonoblot analysis of proteins from lipid-raft- and soluble-membrane fractions probed with anti-human Cav-1 Ab. Lane 1 shows lipid raft proteins isolated from the human HT-29-MDR cell line as a control of caveolin-1 presence (Lavie et al., 1998Go). Lanes 2-7 represent fraction 4, 5 and 6 of the sucrose gradient that contained lipid rafts. Lanes 8 and 9 correspond to the soluble membrane fraction (fraction 11). Caveolin-1 oligomers are indicated by arrowheads. (B) Silver staining of an identical gel. Lane 1 shows protein-size marker. (C) Comparison of caveolin-immunodetected patterns of Sf9 cells (lanes 1 and 3) and human HT-29-MDR cells (lanes 2 and 4). Western blot analysis of total protein extracts incubated with anti-human Cav-1 Ab. Lanes 1 and 2 show caveolin monomers and oligomers of Sf9 cells and HT-29-MDR cells, respectively. Lanes 3 and 4 show that the caveolin bands disappear from the same blot upon probing with the same amount of anti-human Cav-1 Ab that was incubated earlier with an equivalent amount of HT-29-MDR total proteins under the same conditions to pre-absorb the anti Cav-1 Ab.

 

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, 2002Go). 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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Fig. 5. Reduced Cry1C binding to G2-M-arrested Sf9 cells. (A,C) Cry1C (1 µg/ml in A and 2.5 µg/ml in C) was incubated with normal and G2-M-arrested cells for 5 or 90 minutes. After washing off the non-bound toxin, total proteins (9 µg/lane) were separated by SDS-PAGE and Cry1C binding was detected by western analysis using anti-Cry1C Ab. (Lane 1) Cry1C marker, 100 ng. (Lane 2) Sf9 cells with no Cry1C in the binding reaction. (Lane 3) Sf9 cells after 5 minutes incubation with Cry1C). (Lane 4) G2-M cells after 5 minutes incubation with Cry1C. (Lane 5) Sf9 cells after 90 minutes incubation with Cry1C. (Lane 6) G2-M cells after 90 minutes incubation with Cry1C. Quantification of band-intensity (indicated below the corresponding lanes in arbitrary units) showed that G2-M cells bound 10 times less Cry1C than the untreated cells. (B) Blot shown in (A) probed with anti-actin Ab as a loading control.

 
Binding of Cry1C to normal and G2-M-arrested Sf9 cells
Cry1C toxicity, similarly to that of other Cry1 toxins, requires the involvement of specific membrane receptors (Schnepf et al., 1998Go) and other, as yet unknown, membrane components (Avisar et al., 2004Go). To correlate the lack of Cry1C sensitivity during M phase with Cry1C-binding, normal and G2-M cells were exposed to relatively high concentrations of Cry1C (1 or 2.5 µg/ml) to enable the detection of Cry1C-binding to the latter cells with lesser Cry1C sensitivity. Both cell cultures were sensitive to these high toxin concentrations (Fig. 2). However, G2-M cells were found to be less sensitive (Fig. 2) and bound considerably less toxin than the normal cells (Fig. 5). Moreover, Cry1C binding to normal cells was already detected after a 5-minute incubation with Cry1C at the higher concentration (2.5 µg/ml; Fig. 5C, lane 3) and only after a 90-minute incubation with Cry1C at the lower concentration (Fig. 5A and C, lane 5). Binding to the G2/M cells, however, was only detected after a 90-minute incubation with Cry1C at the high concentration of 2.5 µg/ml (Fig. 5A and C, lane 6), although equal amounts of proteins were tested in the binding assays (Fig. 5B). The arrested cells bound 10 times less toxin than normal cells according to quantification of the immunodetected bands in three independent binding experiments. Thus, these data suggest that, during mitosis, Sf9 cells are Cry1C-tolerant owing to a strict reduction in toxin-binding-capacity.


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 References
 
By using Sf9 cells as a model system for analyzing Cry1C-membrane-interactions, we demonstrated that dividing cells loose their Cry1C-sensitivity and therefore tolerate the presence of the toxin throughout the entire M phase and early G1 phase. Their daughter cells regain Cry1C-sensitivity during G1-phase progression. This phenomenon was initially observed in dividing cells of log-phase cultures and further confirmed by arresting the cells at G2-M transition.

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., 2000Go; Drews et al., 1995Go). 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., 2000Go; Villalon et al., 1998Go). Sf9 cells that were kept in isotonic buffers were affected by lower concentrations of Cry1C (Kwa et al., 1998Go), 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, 2002Go; Rajagopal et al., 2003Go; Wachtler et al., 2003Go). 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, 2002Go), 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, 2004Go). 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., 2002Go). 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., 2002Go; Rajagopal et al., 2002Go). These data indirectly suggest the involvement of lipid rafts also in Cry1C–membrane 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., 2003aGo; Abrami et al., 2003bGo).

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., 2003Go; Morin et al., 2003Go), although field resistance of Plutella xylostella was generated by Bt sprays (Shelton et al., 1993Go). 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, 2002Go). Naturally mutated host genes encoding a cadherin-like receptor (Gahan et al., 2001Go; Morin et al., 2003Go) or artificially mutated genes encoding ß-1,3-glycosyltransferase (Griffitts et al., 2001Go) 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, 2002Go; Oppert et al., 1997Go) or to higher gut-proteolytic-activity that might lead to toxin degradation (Loseva et al., 2002Go). 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., 2004Go; Ferre and Van Rie, 2002Go).

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., 2003Go; Loeb et al., 2001Go). 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., 2004Go), intracellular trafficking of proteins, toxins and pathogens that require specific interactions with lipid raft resident proteins and sphingolipids (Helms and Zurzolo, 2004Go; Taieb et al., 2004Go), and many signaling pathways (Zhai et al., 2004Go). 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, 2004Go). 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.


    Acknowledgments
 
We thank M. Lescovitch, O. Erster, N. Jain and D. Ravid for providing the HT29-MDR cells, the know-how and advice for lipid raft isolation, and caveolin identification. We also thank J. Gressel for critically reading the manuscript, and N. Chejanovsky for providing the Sf9 cell line and for advice on how to maintain the cultures. This research was supported by the BSF grant (2001235) from the United States-Israel Binational Science Foundation (BSF), Jerusalem, Israel.


    Footnotes
 
Supplementary material available online at http://jcs.biologists.org/cgi/content/full/118/14/3163/DC1


    References
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 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
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