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Applied and Environmental Microbiology, October 2008, p. 5875-5881, Vol. 74, No. 19
0099-2240/08/$08.00+0     doi:10.1128/AEM.01228-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Protozoal Digestion of Coat-Defective Bacillus subtilis Spores Produces "Rinds" Composed of Insoluble Coat Protein

Alicia Monroe Carroll,^1, Marco Plomp,² Alexander J. Malkin,² and Peter Setlow^1*

Department of Molecular, Microbial and Structural Biology, University of Connecticut Health Center, Farmington, Connecticut 06030,¹ Chemistry, Materials, Earth and Life Sciences Directorate, Lawrence Livermore National Laboratory, Livermore, California 94551²

Received 3 June 2008/ Accepted 28 July 2008

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The Bacillus subtilis spore coat is a multilayer, proteinaceous structure that consists of more than 50 proteins. Located on the surface of the spore, the coat provides resistance to potentially toxic molecules as well as to predation by the protozoan Tetrahymena thermophila. When coat-defective spores are fed to Tetrahymena, the spores are readily digested. However, a residue termed a "rind" that looks like coat material remains. As observed with a phase-contrast microscope, the rinds are spherical or hemispherical structures that appear to be devoid of internal contents. Atomic force microscopy and chemical analyses showed that (i) the rinds are composed of insoluble protein largely derived from both outer and inner spore coat layers, (ii) the amorphous layer of the outer coat is largely responsible for providing spore resistance to protozoal digestion, and (iii) the rinds and intact spores do not contain significant levels of silicon.

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The gram-positive bacterium Bacillus subtilis forms remarkably resistant endospores when it is faced with starvation (36). This process, known as sporulation, produces metabolically dormant spores that are resistant to heat, radiation, and desiccation, as well as to predation by the protozoan Tetrahymena thermophila (17, 26, 36). These spores remain dormant until they sense that nutrients are available, at which time they can undergo germination and outgrowth to change back into growing cells (35).

Several layers are assembled around the developing forespore during sporulation. The outermost layer of spores of most species is the coat, a multilayer structure made of more than 50 proteins in B. subtilis spores (7-9, 13). It is thought that the coat proteins together provide resistance to potentially toxic chemicals, as well as predation, but the coat has limited involvement in resistance to heat, radiation, or mechanical disruption (14, 17, 36). It has been difficult to comprehensively study the proteins of the spore coat, as at least 30% of the total coat protein is insoluble (7, 8). Because proteins can become insoluble when they are covalently cross-linked, it is thought that a number of coat proteins participate in this type of interaction (1, 13). SodA, a superoxide dismutase that may catalyze the formation of dityrosine bridges, and Tgl, a bacterial transglutaminase that forms -(-glutamyl)-lysine isopeptide bonds, have been suggested as proteins that may catalyze cross-link formation in the spore coat (12, 18, 19, 24, 40).

Two main layers of the B. subtilis spore coat have been visualized by electron microscopy. The outer spore coat is thick and layered, while the inner spore coat is composed of several fine lamellae (9). Proper assembly of these layers is dependent on a number of morphogenetic proteins, including SpoIVA, SpoVID, SafA, CotE, CotH, and CotO, as well as the transcription factor GerE, and loss of any one of these proteins alters spore coat assembly, as well as the final coat structure (13). According to the models of spore coat assembly that have been described, during sporulation SpoIVA is produced in the mother cell immediately after asymmetric division and assembles around the forespore surface (34). Once SpoIVA has assembled, a CotE ring, whose formation is SpoIVA dependent, assembles 75 nm from SpoIVA (10, 28, 33). The space between SpoIVA and CotE is called the matrix, and as sporulation continues, the matrix becomes the inner spore coat (10), while the outer spore coat forms around the CotE ring, leaving this protein sandwiched between the two layers once coat assembly is complete (21). While coat proteins can be synthesized in the absence of CotE, the outer coat cannot be assembled (21), and without a properly assembled outer coat the spore is vulnerable to chemicals and lytic enzymes (7, 17, 39).

Although a coat-defective spore is sensitive to protozoal predation, at least part of the spore is resistant, as a residue that resembles the coat remains after protozoal digestion (17). Such residues, called "rinds," appear to be hollow, spherical, or hemispherical structures when they are examined by electron and phase-contrast microscopy (17). In this work, we used atomic force microscopy (AFM) and chemical analyses to probe the structure of wild-type and cotE rinds from B. subtilis spores.

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Strains used and spore preparation.
The B. subtilis strains used in this study are PS832 (wild type), PS3394 (cotE::tet) (17), and PS3760 (cotE::tet sodA::cat tgl::ermC) (17). All of these strains are isogenic with strain PS832, a prototrophic derivative of strain 168.

To prepare spores, strains were grown for 3 h at 37°C in Luria-Bertani (22) medium supplemented with the appropriate antibiotic(s), as follows: tetracycline, 10 mg/liter; chloramphenicol, 5 mg/liter; or erythromycin and lincomycin, 1 and 25 mg/liter, respectively. Cells were spread on 2x SG agar plates (25) without antibiotics and incubated at 37°C for 5 days and at 23°C for 2 days. Spores were harvested and purified by sonication and repeated washing with distilled water as described previously (25). All spore preparations were free (>98%) of vegetative and/or sporulating cells, as well as germinated spores, as determined by phase-contrast microscopy.

Growth of Tetrahymena.
A stock culture of T. thermophila strain CU428.2 was grown at 30°C on SPPA medium containing 250 µg/ml penicillin G, 250 µg/ml streptomycin, and 0.25 µg/ml amphotericin B (11). To prepare cultures for spore feeding, 9.5 ml of fresh SPPA medium was inoculated with 0.5 ml of the stock culture and incubated overnight at 30°C with shaking. Cells were harvested and starved for 2 to 4 h as described previously (17).

Spore decoating and preparation of rinds.
To prepare spore rinds, wild-type strain PS832 spores (10 to 20 mg [dry weight]) were decoated by incubation in 1 to 2 ml of decoating buffer (50 mM Tris-HCl [pH 8.0], 1% sodium dodecyl sulfate [SDS], 8 M urea, 50 mM dithiothreitol, 10 mM EDTA) for 90 min at 37°C. The spores were centrifuged, and the pellet was washed six times with 10 mM Tris-HCl (pH 7.4). Decoated wild-type spores or intact spores of cotE strains (7.5 x 10⁸ spores/ml; final optical density at 600 nm, 7.5) plus starved Tetrahymena cells (2 x 10³ cells/ml) were incubated at 30°C with slow shaking in 12.5 ml of 10 mM Tris-HCl (pH 7.4). Rinds accumulated at the bottom of the tube after 24 h; the supernatant fluid was removed, and the rinds were washed and suspended in an equal volume of 10 mM Tris-HCl (pH 7.4). After shaking at 100 rpm and 30°C for 30 min, the rinds again were allowed to settle at the bottom of the tube; the supernatant fluid was removed, and the rinds were suspended in a small volume of 10 mM Tris-HCl (pH 7.4). These rind preparations were free (>95%) of Tetrahymena but contained 10% intact spores adhering to the rinds based on examination by AFM.

Rind integrity experiments.
Decoated wild-type spores (6 mg [dry weight]), wild-type rinds (3 to 9 mg [dry weight]), and PS3760 (cotE sodA tgl) spores and rinds (6 and 3 mg [dry weight], respectively) were prepared as described above. To test rind strength, spores and rinds were either pulverized with 100 mg of glass beads with a dental amalgamator (Wig-L-Bug) using 20 30-s pulses with 30-s intervals between pulses or sonicated with a microtip probe for 20 s using a Heat Systems-Ultrasonics cell disrupter (model W-220F). Samples were centrifuged, and the pellets were suspended in 90 to 200 µl of 2x SDS-polyacrylamide gel electrophoresis (PAGE) sample buffer (24). Samples were boiled for 5 min before 10% polyacrylamide SDS-PAGE was performed (24), and proteins were visualized by staining with Coomassie blue.

In a second experiment to test rind strength, wild-type rinds (3 mg [dry weight]) were suspended in 0.5 ml water and sonicated for 45 s. The sample was divided and placed into into two tubes containing the same volume of rinds, and one tube was centrifuged. The pellet from the latter tube was suspended in 250 µl of 2x SDS-PAGE sample buffer without bromophenol blue, boiled for 5 min, and centrifuged, and the pellet was suspended in 250 µl of water before sonication for 30 s. Ten microliters of each sample was applied to an agarose-coated microscope slide, a cover glass was applied, the edges of the cover glass were sealed with clear nail polish, and the sample was examined by using differential interference contrast (DIC) microscopy (6).

Amino acid analysis.
PS3394 (cotE) lyophilized rinds (12 mg [dry weight]) were dissolved in 1.2 ml of 70% formic acid, and 10 µl was hydrolyzed for 16 h at 115°C in 100 µl of 6 N HCl-0.2% phenol containing 2 nmol norvalene as a standard. The sample was dried, suspended in 100 µl of 20 mM HCl containing 2 nmol taurine as a second standard, and an amino acid analysis was carried out with a Hitachi model L-8900 amino acid analyzer.

Carbohydrate analysis.
PS3394 (cotE) rinds (9 mg [dry weight]) were washed five times with 1 ml of water, suspended in 3 ml of water, and sonicated for 30 s. Fifty microliters of a sample was placed on a microscope slide and allowed to air dry. Dried slides were prewetted in water and placed in a Coplin jar, and the periodic acid-Schiff (PAS) reagent was used to stain the samples as described previously (2, 20).

Silicon (Si) analysis.
PS3394 (cotE) spores or rinds (17 mg [dry weight]) were suspended in water and centrifuged at 6,000 rpm. The pellet was warmed to 45°C and then mixed with agar on a glass slide heated to 55°C. The slide was cooled to room temperature, and the solidified agar mixture was cut into 1- to 2-mm cubes with a razor blade. The agar cubes were transferred to vials containing a 1:1 mixture of 50% polyvinylpyrrolidone (molecular weight, 10,000) in water and 2.3 M sucrose in 0.1 M sodium phosphate, and the vials were rotated overnight at 4°C. Excess sucrose and polyvinylpyrrolidone were removed, and the samples were frozen in liquid nitrogen. Sections of spore and rind samples were cut at –170°C using a Reichert cryo-ultramicrotome (model Ultracut E), and the sections were collected in homemade platinum loops holding a 1:1 mixture of 2% methylcellulose and 2.3 M sucrose and transferred to copper grids with holey carbon support film backing. The grids were stained by placing them section side down on a solid gelatin surface and incubating them at 45°C for 20 to 30 min to remove the methylcellulose-sucrose solution. The grids were washed with 45°C water, stained on droplets of 4% uranyl acetate (pH 7.0; pH adjusted with oxalic acid) for 5 to 10 min, and placed on drops of a 9:1 mixture of 2% methylcellulose and 4% uranyl acetate at 23°C. The grids were transferred to a fresh solution at 23°C twice and then to drops of a 4°C solution for 5 to 10 min. The grids were picked up in platinum loops, and the solution was wicked off by drawing the edge of the loop along a piece of filter paper. The grids were air dried before they were viewed with a 200-kV FEI Tecnai transmission electron microscope (FEI Company, Hillsboro, OR). Energy-dispersive X-ray spectroscopy (EDS) was used to detect Si in the spores or rinds. EDS data were collected with an accelerating voltage of 200 kV for 100 to 500 s using an EDAX EDS system (EDAX Inc., Mahwah, NJ) and FEI/Emispec TIA software. EDS quantification was based on a standardless analysis.

AFM.
Intact spores, decoated spores, or rinds (3 mg [dry weight]) were suspended in 2 ml of water. Droplets (2.5 µl) of spore or rind suspensions were deposited on plastic coverslips, incubated for 10 min, gently rinsed with double-distilled water, allowed to dry, and transferred to the AFM. Images were collected using a Nanoscope IV AFM (Veeco Instruments, Santa Barbara, CA) operated in tapping mode. For low-resolution analysis of spore samples, fast-scanning AFM probes (DMASP Micro-Actuated; Veeco Instruments) with a force constant of 1 to 5 N/m and a resonance frequency of 200 kHz were utilized. For high-resolution imaging Veeco and NanoWorld etched silicon tips with force constants of 40 N/m and resonance frequencies of 300 kHz were used. Tapping amplitude, phase, and height images were collected simultaneously.

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Spore and rind ultrastructure.
In previous work, the structure of rinds obtained from digested spores was examined using electron microscopy (17). Since AFM has been used to examine the high-resolution structure and assembly of the spore coat of Bacillus and Clostridium species (5, 29-32), AFM was used to compare the structures of rinds and intact spores. This analysis showed that intact wild-type spores were completely or partially covered by a thin amorphous layer without a defined structure (Fig. 1a and b). Directly below the amorphous layer is a rodlet crystalline layer (Fig. 1b), which has parameters similar to those of the B. atrophaeus rodlet spore coat layer (31). In decoated spores the amorphous layer appeared to be largely removed, exposing the rodlet crystalline layer (Fig. 1c). The surface of wild-type rinds appeared to be distinct from that of intact spores, since the amorphous layer was absent in rinds (Fig. 1d to f). A detailed analysis of spore and rind structure is described below.

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FIG. 1. AFM images of wild-type spores and rinds. Intact wild-type and decoated wild-type spores and wild-type rinds were examined by AFM as described in Materials and Methods. (a and b) Intact wild-type spores; (c) decoated wild-type spores; (d to f) wild-type rinds. The arrowheads in panels b, c, and f indicate the rodlet layer. The arrow in panel b indicates the amorphous cover layer, and the arrow in panel f indicates the pitted layer.

Rinds contain insoluble protein.
Rinds are produced after spores with defective coats are digested by Tetrahymena (17). While the origin of the rinds is technically unknown, it is likely that the rinds consist of spore coat protein. To determine if rinds are indeed composed largely of protein, purified rinds from cotE spores prepared in Tetrahymena were acid hydrolyzed and subjected to amino acid analysis. This analysis showed that the rinds contained 60% protein, even when the contamination of rinds with intact spores was taken into account (data not shown).

Given the information on rind protein content, identification of the proteins present in the rinds was attempted by extraction with SDS-PAGE sample buffer, followed by separation by SDS-PAGE and protein identification by mass spectrometry. However, while a significant amount of protein was extracted from pulverized decoated wild-type spores and even from intact wild-type spores, only a miniscule amount of protein was extracted from intact rinds (Fig. 2, lanes 1 and 3, and data not shown). Similarly, only minimal amounts of protein were extracted from pulverized or sonicated wild-type rinds (Fig. 2, lanes 1, 2, and 7). In addition, there was no difference in rind structure before and after boiling and sonication, as determined by phase-contrast or DIC microscopy (data not shown). These results strongly suggest that the proteins in rinds are extremely insoluble. The rinds were, however, broken up by pulverizing them with a dental amalgamator (data not shown), as expected, since even intact spores are completely disrupted by this procedure.

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FIG. 2. Protein extracted from decoated spores and rinds of various strains. Decoated wild-type spores and intact spores of strain PS3760 (cotE sodA tgl) (6 mg [dry weight]) were pulverized, and 9 mg (dry weight) of intact PS3760 spores or decoated wild-type spores were fed to Tetrahymena. Rinds were purified and were sonicated or pulverized as described in Materials and Methods. All samples were suspended and boiled in 2x SDS-PAGE sample buffer before aliquots (2.5% of the total sample) were subjected to SDS-PAGE, although the amounts of samples from rinds that were used were corrected for rind recovery (33%). Lanes 1, 2, and 7, rinds from wild-type spores that were not treated, were pulverized, and were sonicated, respectively; lanes 3 and 6, pulverized wild-type decoated spores and intact PS3760 spores, respectively; lanes 4 and 5, rinds from intact PS3760 spores that were not treated and were pulverized, respectively. Note that lane 7 was taken from a separate gel. The migration positions of molecular mass standards (in kDa) are indicated on the left.

As mentioned above, it has been suggested that much of the insolubility of the spore coat is due to a high degree of protein cross-linking (7-9, 13). Two enzymes that have been suggested to be involved in coat protein cross-linking are SodA and Tgl (12, 18, 19, 24, 40). Therefore, more protein might be extracted from rinds from cotE sodA tgl spores (strain PS3760) that lack an outer coat, as well as Tgl- and SodA-catalyzed cross-links. However, while a large amount of protein was extracted from pulverized strain PS3760 spores (Fig. 2, lane 6), significantly less protein was extracted from rinds of this strain either with or without prior pulverizing (Fig. 2, lanes 4 and 5), although more protein was extracted from these rinds than from wild-type rinds (compare Fig. 2, lanes 1 to 3 and lanes 4 to 6). This indicates that the protein in the mutant rinds is slightly more soluble than the protein in the wild-type rinds. However, again, there was no change in the cotE sodA tgl rind structure after the rinds were boiled in SDS-PAGE sample buffer, as determined by microscopy, and no distinct protein bands were obtained upon SDS-PAGE of extracts of cotE sodA tgl rinds (Fig. 2, lanes 4 and 5).

Since the protein in the cotE sodA tgl rinds was more soluble than the protein in wild-type rinds, partial acid hydrolysis was used to produce peptides that could be analyzed by liquid chromatography-mass spectrometry. Rinds from wild-type and cotE sodA tgl spores were acid hydrolyzed as described in Materials and Methods, except that hydrolysis was performed for only 4 h. Following hydrolysis, samples were analyzed by liquid chromatography-mass spectrometry, and tandem mass spectrometry spectra were searched using the automated Mascot algorithm against the NCBInr database. Several peptides were produced from both types of rinds by partial acid hydrolysis, but remarkably, no B. subtilis proteins were identified (data not shown). These results suggest that the insoluble proteins in the rinds may be part of such a complex, heavily cross-linked network that individual components cannot readily be identified by commonly used techniques. Taken together, these results indicate that while deletion of Tgl and SodA does have an effect on protein solubility, there is likely another cross-linking enzyme(s) in the spore coat.

The insoluble protein in the rinds is coat protein.
Although the rinds were found to contain insoluble protein, it was not clear if this protein originated from the spore coat. Since it was not possible to examine rind structure at the molecular resolution level by phase-contrast or DIC microscopy, AFM was used to visualize the structure of spores and rinds. As noted above, AFM indicated that intact wild-type spores are completely or partially covered by an amorphous layer (Fig. 1a and b). Since such spores have not been decoated, the amorphous layer is most likely the outermost layer of the spore coat. Directly beneath this amorphous layer is a second layer with a rodlet structure (Fig. 1b). It has been shown with spores of several Bacillus species (30) that both external (B. atrophaeus) and internal (B. cereus) layers of the outer spore coat consist of assembled rodlets, which suggests that the rodlets seen in B. subtilis are comprised of outer spore coat protein as well. When wild-type spores were decoated, the amorphous layer was removed, although the rodlet layer remained intact (Fig. 1c). Since Tetrahymena produces rinds from wild-type spores only after spores are decoated, it was expected that at least the amorphous layer would be not be present in the rinds. Indeed, the outer surfaces of all wild-type rinds visualized at high resolution had the rodlet layer (Fig. 1f), indicating that the amorphous layer on rinds had been removed. Note that on some (20 to 40%) wild-type rinds, small patches of pitted and layered inner coat structures were seen (Fig. 1f). These structures were found to assemble beneath the rodlet layer, and each inner coat layer was 6 nm thick (M. Plomp, A. M. Carroll, P. Setlow, and A. J. Malkin, unpublished data). The exposure of these underlying layers may indicate that there was partial digestion of the rodlet layer. These results suggest that the amorphous layer is responsible for providing spore resistance to degradative enzymes located in the phagosomes of Tetrahymena. Furthermore, the thickness of the rinds varied from 30 to 40 nm, which indicates that the rinds are composed of the rodlet and underlying pitted layers (outer coat), as well as several inner coat crystalline layers.

CotE is a major spore coat morphogenetic protein, and in its absence the outer coat does not assemble properly (21). It was expected that in intact cotE spores the amorphous and possibly the rodlet layer would be absent. Indeed, for most (>90%) cotE spores the outermost structure was formed by three to five layers, each of which was 6 nm thick (Fig. 3a). The step edges of these layers showed a clear growth morphology and likely corresponded to the inner coat layers, as is the case for the coats of Clostridium novyi (32) and B. anthracis (M. Plomp and A. J. Malkin, unpublished results) spores. Note that small patches of rodlet structures or groups of several individual rodlets were seen on the majority (>75%) of cotE spores (Fig. 3a). Patches or large regions of a hexagonal crystalline layer (located between the rodlet layer and the inner coat multilayer structure) covering the spore have also been observed (Fig. 3a and data not shown). Furthermore, some (<10%) cotE spores were completely covered with the rodlet layer and patches of the amorphous cover layer (M. Plomp, A. M. Carroll, P. Setlow, and A. J. Malkin, unpublished data). The presence of the rodlet and hexagonal crystalline layers in some cotE spores suggests that the outer coat may be partially assembled in a small fraction of the cotE spore population, which may explain the presence of intact cotE spores in rind samples (Fig. 3b). AFM images of cotE rinds confirmed the results obtained with intact cotE spores. The outermost layers of the vast majority (>90%) of cotE rinds were formed by the inner coat layers, and small patches of rodlet structures were occasionally seen at the top of the layered structure (Fig. 3c and d). These results suggest that cotE rinds are composed of inner coat proteins and trace amounts of outer coat proteins (the proteins in the rodlet layer). Taken together, these results strongly suggest that the insoluble protein in both wild-type and cotE rinds originates from the spore coat. In addition, these results indicate that the insoluble fraction of the spore coat is composed largely of the proteins in the outer coat (rodlet layer), as well as the proteins from the inner coat layers (coat growth layers).

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FIG. 3. AFM images of cotE spores and rinds. Intact cotE (PS3394) spores and cotE rinds were examined by AFM as described in Materials and Methods. (a) Intact cotE spores; (b to d) cotE rinds. In panel a, the black arrowhead indicates rodlets, the white arrowhead indicates the hexagonal crystalline layer, and the arrow indicates the inner coat growth layers. In panel d, the arrow indicates inner coat growth layers.

Spore rinds are devoid of carbohydrate and silicon.
Although the rinds consist largely of spore coat protein, rinds were also tested to determine whether two other possible constituents, carbohydrate and Si, were present. For carbohydrate analysis cotE rinds were stained for carbohydrate using the PAS reagent. While this reagent does not detect all carbohydrates, it detects carbohydrates containing a 1,2-glycol, including polysaccharides, glycoproteins, and glycolipids. However, the cotE rinds did not stain with the PAS reagent, while a positive control (a section of mouse colon) stained bright magenta, as expected (data not shown).

In addition to the carbohydrate anhalysis, the rinds were also examined to determine whether Si was present. Although Si has not been detected in spores of some Bacillus species (in particular, B. coagulans), Si has been found in the coat and cortex regions of B. cereus and B. megaterium spores (15, 27, 37, 38). The presence of Si in spores (specifically, spores of B. anthracis) is particularly interesting because coating spores with silica facilitates spore dispersion through the air and can "weaponize" B. anthracis spores (23). Thus, if Si is normally absent from spores, its presence in a spore sample might suggest that the spores were weaponized. In addition, Si is found in outer skeletons of a number of types of microorganisms (4) and thus could theoretically be important in rind structure.

EDS was used for elemental analysis of copper grids with holey carbon support film backing containing carbon (C) and Si, and two Si peaks occurred at 1.739 and 1.829 keV along with a large C peak, two small copper (Cu) peaks, and an oxygen (O) peak (Fig. 4A). The oxygen peak was due to copper oxide formed when copper in the grid was exposed to air. EDS has a theoretical Si detection limit of 0.1 to 0.2% (based on the manufacturer's analysis); therefore, at least 1,000 ppm Si (0.1% of the total mass) must be present before Si is detected by EDS. To obtain a background spectrum, a blank copper grid with holey carbon support film backing was analyzed, and it lacked Si but contained C, Cu, and O (Fig. 4A and B). EDS analysis of intact cotE spores showed that there was no Si peak, but the large C peak, the O peak, and the small Cu peaks observed in the carbon grid were present, as were several small uranium (U) peaks (Fig. 4C). The presence of uranium in the sample was due to the uranyl acetate used to stain the spores. cotE rinds gave results similar to those obtained for intact cotE spores; there was no Si peak, but the large C peak, the O peak, and several small Cu and U peaks were present (Fig. 4D). These results indicate that there is no detectable level of Si (0.2% of the total mass) in either intact cotE spores or cotE rinds. However, since cotE spores do not have the amorphous layer of the spore coat, it is possible that this region could contain Si.

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FIG. 4. EDS analysis of cotE spores and rinds. cotE (PS3394) rinds were examined by EDS as described in Materials and Methods. (A) Blank copper grid with support film backing containing carbon and silicon; (B) blank copper grid with holey carbon support film backing containing no silicon; (C) intact cotE spores; (D) cotE rinds. The box in each panel indicates the area that was analyzed. In the spectra C indicates carbon, O indicates oxygen, Cu indicates copper, Si indicates silicon, and U indicates uranium. Note that in panels C and D the carbon peak is not labeled but is in the same position as it is in panels A and B.

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AFM has been used previously to examine the ultrastructure of spores of several Bacillus species, as well as spores of C. novyi (5, 29, 30, 32). Here, this high-resolution technique was used to examine the spore coat of B. subtilis. Our results indicate that the spore coat consists of several layers, including an amorphous cover layer, a rodlet layer, a hexagonal crystalline layer, and a multilayer inner coat. Based on an analysis of the layers of the spore coat that are present in spores and rinds of various strains, we concluded that (i) rinds produced after protozoal digestion of B. subtilis coat-defective spores are composed of insoluble protein that originates from the rodlet layer of the outer spore coat, as well as the inner coat layers, and (ii) the amorphous outermost layer of the outer spore coat is responsible for resistance to protozoal predation.

Previous work in our laboratory has shown that deletion of a major coat morphogenetic protein (CotE, SafA, or SpoVID) results in the production of rinds after spores are fed to Tetrahymena (17). As mentioned above, spores that lack CotE can synthesize the outer spore coat but cannot assemble it (21). In the absence of SafA, spores produce a coat whose structure and composition are slightly perturbed (8, 16). spoVID spores show a mixed phenotype; some spores have a coat that is mislocalized as swirls in the mother cell cytoplasm, while other spores have a coat that is properly localized but not fully attached to the forespore (3). Here, we show that only when they lack the amorphous outer layer are spores digested to rinds by Tetrahymena. Taken together, these results suggest that this outer layer of the outer spore coat is also absent from safA spores and is absent from or not fully attached to spoVID spores. Interestingly, these spores are also sensitive to treatment with lysozyme (3, 17), which indicates that the amorphous layer likely plays a role in spore resistance to lytic enzymes as well. It is important to perform AFM studies with safA and spoVID spores to determine the structure of their spore coat, and this work is in progress.

In contrast to cotE, safA, and spoVID spores, sodA, tgl, or sodA tgl spores are not sensitive to protozoal predation (17). Presumably, the amorphous cover layer is intact in sodA tgl spores, suggesting either that there are no SodA- or Tgl-catalyzed cross-links in this layer of the spore coat or that the cross-links do not contribute significantly to spore resistance to lysozyme or protozoal predation. Since the outer amorphous layer is readily removed during decoating, it appears likely that this layer has minimal, if any, cross-links. Cross-links are more likely present in the inner insoluble fraction of the spore coat that remains attached to the spore during decoating. However, the structure and integrity of rinds from cotE sodA tgl spores, in which the amorphous layer and the two candidate cross-linking enzymes were absent, were not significantly different from the structure and integrity of wild-type rinds. This result and the fact that the analysis of peptides generated by partial acid hydrolysis of rinds of cotE sodA tgl spores could not identify B. subtilis proteins suggest that the great majority of protein in the rinds is still cross-linked, and thus there must be at least one additional cross-linking enzyme in spore coats.

Given that so much of the spore coat protein is insoluble and presumably highly cross-linked, the proteins must be held together tightly in this rind structure for a purpose. One possibility is that they play a role in resistance to mechanical disruption (14), and this is consistent with the resistance of rinds to sonication and boiling in SDS. Another possibility is that the insoluble coat proteins contribute to spore resistance to potentially toxic chemicals. While cotE spores are generally much more sensitive to many toxic chemicals than intact spores (24), these spores with defective coats are still much more resistant to such agents than growing cells are. Perhaps it is the insoluble protein meshwork that is responsible for the chemical resistance of cotE spores.

One final notable observation is the absence of Si in cotE spores and rinds. As mentioned above, Si has been implicated in the weaponization of B. anthracis spores (23); however, to our knowledge, B. anthracis spores have never been tested to determine whether Si is present. Since B. subtilis is often used as a model organism for B. anthracis, it seems likely that Si is not present in spores of this species as well. Although Si was not found in the spore or rind samples, various amounts of carbon, oxygen, and copper were detected. Since cotE spores and rinds are composed of carbon- and oxygen-containing molecules, it was expected that the levels of these elements would increase in these samples. The amount of copper in the samples differed for two possible reasons. First, the amount of copper scattering during EDS analysis increased as the sample approached the edge of the grid. The cotE spore sample was located closer to the edge of the grid than the cotE rind sample. Second, the height of each peak in the spectrum was dependent upon the duration of EDS data collection, which varied from 100 to 500 s in our experiments and may have contributed to the differences in peak height in the samples. Importantly, we show here that EDS can provide a relatively simple and useful method for detecting elements in spores and could therefore be used to determine whether B. anthracis spores naturally contain Si.

 
We are grateful to Ann Cowan, Larry Klobutcher, and Nancy Ryan (University of Connecticut Health Center) for assistance with DIC microscopy, preparation of Tetrahymena cultures, and PAS staining. We also thank Myron Crawford and Fernando Pineda at the W. M. Keck Facility at Yale University for amino acid analysis and Kurt Langworthy and John Donovan at the CAMCOR NanoFab Facility at the University of Oregon for EDS analysis.

This work was supported by a grant GM19698 from the NIH to P.S. Part of this work was performed under the auspices of the U.S. Department of Energy by Lawrence Livermore National Laboratory under contract DE-AC52-07NA27344.

 
* Corresponding author. Mailing address: Department of Molecular, Microbial and Structural Biology, University of Connecticut Health Center, Farmington, CT 06032. Phone: (860) 679-2607. Fax: (860) 679-3408. E-mail: setlow{at}nso2.uchc.edu

Published ahead of print on 8 August 2008.

Formerly Alicia Monroe.

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Applied and Environmental Microbiology, October 2008, p. 5875-5881, Vol. 74, No. 19
0099-2240/08/$08.00+0     doi:10.1128/AEM.01228-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

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