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Accepted Article
Extracellular Vesicles Produced by the Gram-positive Bacterium Bacillus subtilis are Disrupted
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by the Lipopeptide Surfactin
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Running Title: Vesicle Production and Disruption in Bacillus subtilis
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Lisa Brown1*, Anne Kessler1, Pablo Cabezas-Sanchez3, Jose L. Luque-Garcia3, Arturo
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Casadevall1,2
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1
Department of Microbiology and Immunology, Albert Einstein College of Medicine, Bronx, NY
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10461, USA
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2
Department of Medicine, Albert Einstein College of Medicine, Bronx, NY 10461, USA
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3
Department of Analytical Chemistry, Universidad Complutense de Madrid, Madrid, Spain.
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*For correspondence E-mail: [email protected]; Tel: (+1) 718 430 3730; Fax:
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(+1) 718 430 8701.
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This article has been accepted for publication and undergone full peer review but has not been through the copyediting, typesetting, pagination and proofreading process, which may lead to differences between this version and the Version of Record. Please cite this article as doi: 10.1111/mmi.12650 This article is protected by copyright. All rights reserved.
Accepted Article
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Summary/Abstract
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Previously, extracellular vesicle production in Gram-positive bacteria was dismissed due
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to the absence of an outer membrane, where Gram-negative vesicles originate, and the difficulty
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in envisioning how such a process could occur through the cell wall. However, recent work has
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shown that Gram-positive bacteria produce extracellular vesicles and that the vesicles are
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biologically active. In this study, we show that Bacillus subtilis produces extracellular vesicles
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similar in size and morphology to other bacteria, characterized vesicles using a variety of
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techniques, provide evidence that these vesicles are actively produced by cells, show differences
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in vesicle production between strains, and identified a mechanism for such differences based on
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vesicle disruption. We found that in wild strains of B. subtilis, surfactin disrupted vesicles while
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in laboratory strains harboring a mutation in the gene sfp, vesicles accumulated in the culture
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supernatant.
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anthracis, indicating a mechanism that crossed species boundaries. To our knowledge, this is the
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first time a gene and a mechanism has been identified in the active disruption of extracellular
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vesicles and subsequent release of vesicular cargo in Gram-positive bacteria. We also identify a
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new mechanism of action for surfactin.
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Introduction
Surfactin not only lysed B. subtilis vesicles, but also vesicles from Bacillus
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In recent years, several laboratories have reported that bacteria, fungi, and parasites
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produced extracellular vesicles, which in some instances contain proteins associated with
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virulence.
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focused on the outer membrane vesicles (OMV) produced by Gram-negative bacteria (Kuehn;
The most extensive studies of extracellular vesicle production in bacteria have
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Kesty, 2005; Ellis; Kuehn, 2010; Kulp; Kuehn, 2010).
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concentrated and protected manner, and therefore are potent virulence factors of pathogenic
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Gram-negative bacteria. These vesicles contain toxins, DNA, immunomodulatory compounds,
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communication factors, and adhesins and have been associated with cytotoxicity, bacterial
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attachment, intercellular DNA transfer, and invasion (Dorward et al., 1989; Kuehn; Kesty, 2005;
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Ellis; Kuehn, 2010; Maldonado et al., 2011). Until recently, vesicle production by Gram-
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positive bacteria was overlooked due to the inference that differences in the cell wall structure of
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Gram-negative and Gram-positive bacteria precluded their release.
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bacteria, the cell wall of Gram-positive bacteria consists of primarily peptidoglycan and does not
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have an outer membrane. However, the discovery of extracellular vesicles in fungi, which also
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have large cell walls, catalyzed the search of vesicular structures in Gram-positive bacteria
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(Rodrigues et al., 2007).
Accepted Article
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OMVs can release materials in a
Unlike Gram-negative
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The first evidence for vesicles in Gram-positive bacterial cells was provided by Dorward
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and Garon but they were unable to establish function (Dorward; Garon, 1990). More recently,
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vesicle production was described in Mycobacterium ulcerans, Staphylococcus aureus,
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Mycobacterium tuberculosis, and Bacillus anthracis (Marsollier et al., 2007; Lee et al., 2009;
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Rivera et al., 2010; Gurung et al., 2011; Hong et al., 2011; Prados-Rosales et al., 2011; Thay et
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al., 2013). M. tuberculosis vesicles contain compounds that modulate TLR2 responses while
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those of B. anthracis contain anthrax toxin components, indicating that Gram-positive membrane
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vesicles are potentially involved in pathogenesis (Rivera et al., 2010; Prados-Rosales et al.,
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2011). This observation suggested that Gram-positive cells, like Gram-negative bacteria, utilized
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vesicle secretion to deliver concentrated amounts of toxins and other compounds into the
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extracellular space (Marsollier et al., 2007; Lee et al., 2009; Rivera et al., 2010; Prados-Rosales
This article is protected by copyright. All rights reserved.
et al., 2011; Hong et al., 2011; Gurung et al., 2011; Thay et al., 2013). The disruption of B.
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anthracis vesicles by serum was identified as a potential mechanism for vesicle cargo release, as
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it is important to understand what happens to vesicles and their contents after release from the
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cell (Wolf et al., 2012). Recently, vesicle production has also been described for Streptococcus
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pneumonia and pneumococcal vesicles are used to release pneumolysin (Olaya-Abril et al.,
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2014).
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In this work, we explored vesicle production in the Gram-positive, endospore-forming
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bacterium, Bacillus subtilis. B. subtilis is considered the model system for Gram-positive
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bacteria and is therefore an excellent organism to study the genetic and cellular machinery
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involved in Gram-positive vesiculogenesis. Two common strains of B. subtilis studied were the
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3610 environmental strain, which forms robust biofilms and the laboratory strain 168, which has
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attenuated biofilm production. Both strains produced varying amounts of recoverable vesicles
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under different conditions. Environmental strains of B. subtilis are often not easily genetically
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manipulated; therefore domesticated lab strain 168 was created to be easily transformable and
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used for genetic studies in Gram-positive bacteria (Earl et al., 2007). It is hypothesized that both
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strains 168 and 3610 are descendants of B. subtilis Marburg strain and vary from each other by
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only 22 SNPs (Zeigler et al.,2008). These limited genetic differences and ability to isolate
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differing amounts of vesicles make these two strains ideal to study the mechanism of vesicle
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production in B. subtilis. McLoon et al identified five of these genetic differences between
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strains 168 and 3610 that are involved in biofilm formation (mutations in 168 in the genes sfp,
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epsC, swrA, PdegQ, and lack of the plasmid-borne gene rapP) (McLoon et al., 2011). Further
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study of these two strains has led us to identify the gene, sfp, which affects the amount of
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vesicles recovered in B. subtilis. Lab strains, including strain 168, harbor a mutation in sfp
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resulting in a truncated, nonfunctional protein, whereas environmental strains, like strain 3610,
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harbor a wild type copy of this gene (Nakano et al., 1988).
Accepted Article
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Sfp is a 4’-phosphopantetheinyl transferase which transfers a phosphopantetheine group
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from coenzyme A to a peptidyl carrier protein during biosynthesis of the lipopeptide, surfactin,
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and the siderophore, bacillibactin, in B. subtilis (Nakano et al., 1988; Quadri et al., 1998; May et
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al., 2001). We found that strains harboring a functional sfp gene produced very few recoverable
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vesicles whereas strains with nonfunctional copies of sfp, produced a massive amount of
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recoverable vesicles. To determine the mechanism behind this difference in vesicle quantity, and
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how sfp was affecting it, we explored the effect of surfactin on vesicles. We found that vesicles
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from B. subtilis and B. anthracis were indeed disrupted by the presence of surfactin. And this
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was not surprising considering surfactin can perturb lipid membranes (Bernheimer; Avigad,
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1970; Heerklotz; Seelig, 2001; Carrillo et al., 2003). This data suggests that the production of
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surfactin serves as a mechanism for vesicular cargo release in B. subtilis, as well as the ability to
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disrupt vesicles from B. anthracis, indicating a mechanism that reaches across species barriers.
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This work sets the stage for dissecting the function of vesiculogenesis in a genetically tractable
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Gram-positive bacterium.
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Results
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Bacillus subtilis Produces Extracellular Vesicles
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We recovered vesicles from the culture supernatants of Bacillus subtilis lab strain 168,
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wild type strain 3610, AM373, RL3090, AD3610, SL3610, and RL2663 (Table 1).
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morphology and color of the vesicle pellet from each strain was the first indication that there
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were differences in vesicle preparations among strains. Centrifugation of strain 168 culture
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The
supernatants produced a large, reddish-brown vesicle pellet whereas centrifugation of strain 3610
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culture supernatant and 168 heat-killed preparations produced only small, clear pellets (Fig. 1A-
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C). A massive amount of vesicles from strain 168 were visualized by both negative and positive
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staining transmission electron microscopy (TEM) (Figure 1D-E). In contrast, very few vesicles
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were identified in electron micrographs of identical preparations of vesicles from strain 3610.
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Due to the abundance of vesicles in the supernatant of strain 168, most of the subsequent
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characterization was performed on vesicles isolated from this strain.
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Given the propensity of lipids to organize into vesicle-like structures, considerable effort
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was devoted to establishing that recovered vesicles were not artifacts of broken cells or a result
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of self-assembly by extracellular lipids. Consistent with a requirement for cell viability in vesicle
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production, we observed very few vesicles by negative staining TEM when heat-killed 168 cells
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were added to bacterial media, allowed to incubate at 37o C for 18 h in a shaker, and then
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processed for vesicle preparation under the same protocol (data not shown).
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To exclude the possibility that the recovered vesicles were the result of spontaneous
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extracellular assembly of secreted lipids we performed an experiment where the fungal
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polysaccharide glucuronoxylomannan (GXM), of Cryptococcus neoformans, was added to the
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bacterial culture and then its presence and absence inside vesicles was measured by immunogold.
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To ensure that the GXM was small enough in size to become encapsulated inside vesicles it was
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fractionated to less than 10 kDa in size by ultrafiltration through selective molecular mass
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membranes. This size retains reactivity with specific monoclonal antibodies (mAbs). We
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reasoned that if vesicles were forming spontaneously from lipids, they would contain far more
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GXM than vesicles assembled by the bacterial cell (for a scheme of the experimental design see
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Figure 2A), and that was precisely what was measured. As a control, we sonicated vesicles
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isolated from cultures incubated with GXM, expecting that these would contain more GXM,
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since subsequent self-assembly in the presence of GXM would result in trapping of the
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polysaccharide in newly formed vesicles. TEM images of the natural vesicle preparations
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showed the presence of GXM in the sample (Figure 2B).
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significantly more GXM was inside vesicles that reformed after sonication compared to natural
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vesicles produced from live bacterial cultures (6.9% and 2.9%, respectively) (Figure 2C). These
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results provided strong support for the notion that vesicles are produced by the cell and are not
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the result of spontaneous organization of lipids in the media. Sonicated vesicle diameter was also
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significantly smaller and varied less than that of natural vesicles, indicating another difference
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between B. subtilis-produced vesicles and those that reformed from the aggregation of lipids
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(94.54 ± 1.869 and 115.4 ± 2.69) (Figure 2D). We conclude that B. subtilis, like other Gram-
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positive bacteria, actively produce extracellular vesicles (Marsollier et al., 2007; Lee et al., 2009;
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Rivera et al., 2010; Prados-Rosales et al., 2011).
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Physical Characterization of B. subtilis Vesicles
Accepted Article
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In accordance to predictions,
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Vesicle preparations from planktonic B. subtilis cultures were studied using dynamic
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light scattering (DLS) to determine the approximate size of the vesicles. DLS revealed two
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populations of approximately 50 and 150-250 nm in preparations from strain 168 (Figure 3B).
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This size distribution is similar to that reported for vesicles produced by other bacteria (Kuehn;
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Kesty, 2005). DLS analysis of vesicle preparations from B. subtilis strain 3610 under the same
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conditions revealed less distinct distributions and the presence of larger structures, which may be
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flagella given their abundance in TEM micrographs.
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Analysis of strain 168 supernatant pellets by TEM with negative staining revealed large
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numbers of vesicles (Figure 1D) whereas very few vesicles were recovered from strain 3610
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when isolated from the same culture conditions. Images of strain 168 vesicles obtained by
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embedded TEM were measured using ImageJ software, and the result was a mean diameter of
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137.7 nm (Figure 1E). However, because vesicles are spherical, and TEM represents transverse
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sections, a 1.27 correction factor for these 2D measurements was utilized as suggested by Kong
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et al to estimate the actual equatorial dimensions (Kong et al., 2005). With the 1.27 correction
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factor, the mean diameter of vesicles was 174.4 nm (Figure 3A). It is noteworthy that when DLS
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and TEM measurements are compared, TEM tends to underestimate size while DLS
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overestimates the size of samples (Egelhaaf et al., 1996; Hong et al., 2011). TEM images of
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strain 168 vesicles indicated that some vesicles were more electron dense than others. Smaller
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vesicles were significantly more electron dense than larger vesicles (Figure 3C).
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diameters were categorized visually in three groups based on electron density with mean
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diameters of 214.5 nm for least dense (+), 152.2 nm for medium dense (++), and 120.4 nm for
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most dense (+++) (Fig. 3D-F).
Accepted Article
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Vesicle
168
Spherical protrusions from bacterial cells with dimensions similar to vesicles were
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observed in electron micrographs of B. subtilis cells. TEM images of strain 168 cells showed
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rounded structures strongly suggestive for putative vesiculation in the cell membrane (Figure
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4A-B).
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staining TEM in various stages of release (Figure 4C). Finally, SEM images of strain 168 cells
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also showed multiple protrusions on cells consistent with the emergence of vesicles (Figure 4D).
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Although we recovered various quantities of vesicles from strains 168 and 3610, it is notable that
Structures similar to vesicles were also identified on strain 168 cells by negative
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the number of putative vesicle protrusions on cells was not significantly different (data not
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shown).
Accepted Article
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Bacterial cells were labeled with
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C palmitic acid (14C PA), which is incorporated into
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vesicles, to study stability, production over time, and to compare recoverable vesicle quantity
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among strains (Rivera et al., 2010; Prados-Rosales et al., 2011; Wolf et al., 2012). B. subtilis
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strain 168 vesicles were isolated after 0, 4, 12, 18, and 24 h of growth and radioactivity of the
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vesicle pellets were analyzed as fractions of the total culture. Strain 168 manifested increased
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vesicle production over time, which leveled off after 12 h of growth (Figure 5A). Stability of
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strain 168 vesicles in PBS was also measured by
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incubated for 0, 24, 48 and 72 h before being spun down, counted and the ratio of vesicle versus
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supernatant radioactivity was analyzed. Vesicle pellets contained high levels of radioactivity for
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24 h but subsequently decreased, indicating that vesicles were stable for approximately 24 h
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(Figure 5B). DLS measurements of vesicle stability preparations from each time point showed
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consistent peaks of approximately 50 and 100-150 nm in diameter (Figure 5C-F).
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Chemical Composition of Vesicles
14
C PA labeling. Vesicle preparations were
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Proteomic analysis was performed on strain 168 vesicles, cell membrane fractions, and
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cells. B. subtilis 168 vesicles were enriched in 30 proteins and shared 46 with both the cell and
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cell membrane fractions (Figure 6A). Vesicles were enriched in lipoproteins and contained many
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siderophore-binding proteins (Table S2). SunI, the sublancin immunity protein, was found
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associated with the vesicles (Dubois et al., 2009). Most of the proteins in the vesicle preparation
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were membrane proteins, suggesting that the vesicles originated from the membrane of the
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bacterial cell (Table S2). SDS-PAGE gel of vesicle preparations from strains 168, 3610, and
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RL2663 exhibited similar profiles with only a few differing bands (Figure 6B).
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Vesicle Membrane Potential
Accepted Article
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The ζ potential of strain 168 vesicle preparations were determined to be -54.2917 +/- 9.24
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and -36.724 +/- 10.2 in 10 mM Dextrose and 1 mM KCl, respectively (Figure 6C).
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Vesicles are Constituents of Biofilms
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Negative staining TEM revealed the presence of vesicle-like structures in B. subtilis
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biofilms. After 4 and 8 d of growth, vesicles were isolated from disrupted B. subtilis strains 168
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(i-ii) and 3610 (iii-iv) biofilm pellicles. Despite the previously mentioned differences in both
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recovered vesicle quantity and biofilm growth between the two strains, similar amounts of
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vesicles were isolated from all cultures and visualized using electron microscopy with negative
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staining (Figure 7A). The size distribution of the vesicles isolated from strains 168 and 3610
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biofilms was determined by DLS, which revealed the presence of two distinct size populations
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across all biofilm vesicle samples with measurements comparable to those described for
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supernatant vesicles from planktonic cultures (Figure 7B).
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Samples of intact B. subtilis strains 168 and 3610 biofilm were further visualized using
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TEM and SEM. Embedded biofilm TEM images of strain 3610 biofilm showed vesicles as part
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of the biofilm matrix (Figure 7C). For SEM, biofilm samples were prepared with ferrocyanide in
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a process that ultimately peels back the outer layer of the biofilm to allow for viewing of the
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internal biofilm constituents. SEM images generated from the ferrocyanide-treated samples
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indicated the presence of putative vesicular events on cells within the B. subtilis strain 3610
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biofilm (Figure 7D). This article is protected by copyright. All rights reserved.
Vesicle Quantification
Accepted Article
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A dramatic difference in vesicle quantity between strains 168 and 3610 was apparent in
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TEM micrographs. To further explore this difference, we utilized metabolic labeling to gain
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more quantifiable information about vesicles from various strains. Cells from strains 168 and
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3610 were labeled with 14C PA and then we measured radioactive counts in cells, flow through
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(FT), supernatant, and vesicle pellets, with each measurement determined relative to the total
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sample (Figure 8A-B). Strain 168 had significantly more radioactivity incorporated into the
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vesicle pellet than strain 3610, even though both strains took up a similar amount of radioactivity
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in the initial labeling step (Figure 8C). This data, along with the DLS and electron micrographs,
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provides further support for the observation that the laboratory strain 168 produces more
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recoverable vesicles than environmental strain 3610.
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Metabolic labeling and density gradient centrifugation was utilized to compare vesicle
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quantity and density among various B. subtilis strains (Table 1). Cultures of each strain were
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pulsed with
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labeled samples were also recovered and counted; all fraction radioactivities are represented as a
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percentage of the whole sample.
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C PA and vesicles were isolated. Cells, FT, and supernatant fractions of the
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Strains 168 and AD3610 showed a significantly higher percentage of radioactivity levels
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in the vesicle pellet than pellets of strains 3610, SL3610, and RL3090 (Figure 8D). This indicates
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that supernatants from strains 168 and AD3610 contain a larger quantity of recoverable vesicles
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than 3610, SL3610, and RL3090. Strains 168 and AD3610 have a nonfunctional sfp gene,
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whereas strains 3610, SL3610, and RL3090 harbor fully functional copies of the gene.
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The vesicle preparations were subjected to an Optiprep density gradient and subsequently
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0.5 ml fractions were recovered (Figure 8E). Fractions containing the majority of radioactivity
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were 6, 7, and 8, and the presence of vesicles in these fractions was confirmed by TEM. Outer
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membrane vesicles from Pseudomonas aeruginosa were also found in similar fractions (Tashiro
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et al., 2010).
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Disruption of Vesicles by Surfactin
Accepted Article
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We found that we were able to recover large amounts of vesicles from strains that were
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unable to produce surfactin due to a mutation in sfp and very few vesicles from strains with a
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functional sfp gene. This association suggested to us the possibility that vesicle quantities and
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surfactin expression were linked. The gene, sfp, is involved in the biosynthesis of the lipopeptide
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antibiotic, surfactin, in B. subtilis. Because surfactin is known to disrupt membranes, we sought
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to determine if the presence of surfactin in these sfp+ strains resulted in lower amounts of
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recoverable vesicles. Strain RL2663 (3610 background) has a functional sfp gene but a mutation
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in srfAA, an important module of surfactin biosynthesis, rendering the strain unable to produce
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surfactin (Kearns; Losick, 2003). We were able to recover similar amounts of vesicle pellet
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radioactivity from strain RL2663 as strain 168 (Figure 8F), showing the same phenotype for two
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mutations in the surfactin pathway. We also confirmed the large amount of vesicles by TEM
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(data not shown). This result suggests that the presence of surfactin disrupts vesicles, leading to
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less recoverable reactivity in the vesicle pellet fraction.
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To ascertain whether surfactin action was indeed the mechanism behind the differences in 14
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recoverable vesicles among strains, we incubated
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168 in BHI media, strain 168 cell-free supernatant, strain 3610 cell-free supernatant, PBS, 10%
C PA–labeled vesicles isolated from strain
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EtOH, 10 μg ml-1 surfactin, and 100 μg ml-1 surfactin overnight at 37o C on a nutator and then
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recovered the vesicle pellet and spent supernatants.
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radioactivity was significantly reduced when incubated in supernatant from strain 3610 (surfactin
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competent) but stable in strain 168 supernatant (surfactin deficient) (Figure 9A). We also
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recovered less radioactivity in the pellets of vesicles incubated with purified surfactin and the
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data showed a dose response (Figure 9B). These results indicate that the presence of surfactin
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de-stabilizes vesicles and is responsible for the inability to recover large amounts of vesicles
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from surfactin-producing strains of B. subtilis.
Accepted Article
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We found that recoverable vesicle
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To explore whether the mechanism of vesicle disruption by surfactin extended across
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bacterial species we studied the effect of surfactin on vesicles isolated from B. anthracis. We
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found that pellet radioactivity of
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after incubation with strain 3610 supernatant and purified surfactin (Figure 9C-D). It is notable
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that vesicles from B. anthracis are much less stable under these conditions when compared to B.
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subtilis. B. anthracis vesicle instability was also reported previously (Wolf et al., 2012). We also
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performed the reverse experiments, incubating strain 168 in week-old B. anthracis Sterne,
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Pasteur, and DeltaT cell-free supernatants and did not see a significant change in pellet
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radioactivity compared to BHI and strain 168 supernatant controls (data not shown). This
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indicates B. anthracis does not actively produce a vesicle destabilizing molecule. This result
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extends the role of surfactin to disruption of vesicles from other bacterial species.
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Discussion
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14
C PA–labeled B. anthracis vesicles significantly decreased
The production of extracellular vesicles appears to be a common phenomenon in both
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Gram-positive and Gram-negative bacteria.
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vesicles, the quantity of vesicles varies with the strain, that the gene, sfp, influences vesicle
Here, we established that B. subtilis produces
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recovery through the effect of its product surfactin, and surfactin disrupts vesicles produced by
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different species, suggesting a role for unloading vesicles. These results establish B. subtilis as a
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system for studying the physiology of Gram-positive vesicular production.
Accepted Article
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Any new description of vesiculogenesis by microorganisms must rule out the possibility
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that such vesicles are artifacts of lipid self-assembly and/or damaged membranes resealing into
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spherical structures. Consequently, we devoted considerable effort to establishing that natural
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vesicles were indeed produced from living cells.
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membrane derived vesicles from several lines of evidence: 1) vesicles were recovered from live
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and not dead cells; 2) metabolic labeling with
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radioactivity into vesicle preparations; 3) the vesicles described for B. subtilis had similar
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dimensions to those produced by other microorganisms; 4) vesicle preparations manifested
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similarities and differences in protein content to membranes consistent with both a common
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origin and a separate synthetic pathway; 5) electron microscopy revealed protrusions from cells
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with dimensions comparable to those of extracellular vesicles suggesting that both are the same;
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6) differences in vesicle recovery among B. subtilis strains implied strain-related and -regulated
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differences in vesicle fate; 7) significantly reduced incorporation of GXM in naturally produced
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vesicles (Rodrigues et al.,2007; Marsollier et al.,2007; Lee et al.,2009; Rivera et al.,2010;
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Prados-Rosales et al.,2011; Maldonado et al.,2011) . The latter experiment was designed to
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show that these vesicles were not forming spontaneously in media, which was assessed by taking
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advantage of a fungal polysaccharide for which mAbs suitable for immunogold labeling of
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vesicle preparations are available. We predicted that if vesicles were forming spontaneously by
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the self-assembly of free lipids that we would find a similarly large amount of vesicles
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containing gold particles in both natural and sonicated vesicles. This, however, was not the case,
14
We conclude that B. subtilis produces
C PA
resulted in the incorporation of
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as sonicated vesicles contained significantly more GXM encapsulated within the vesicles.
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Additionally, these structures were smaller and less variable in size than natural vesicles
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recovered from bacterial supernatants. Furthermore, we noted diameter differences between
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sonicated vesicles that presumably had the opportunity to reassemble and naturally recovered
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vesicles, providing further evidence against our vesicle preparations being artifacts of the
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propensity of lipids for self-assembly in solution.
Accepted Article
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Vesicles were isolated from both planktonic cultures and bacterial biofilms. Biofilm is
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considered the primary state in which bacteria exist in the environment therefore it is perhaps not
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surprising that we identified extracellular vesicles within the matrix of B. subtilis biofilm.
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Vesicles isolated from all biofilm preparations were similar in size to planktonic vesicles,
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implying natural vesicle production by B. subtilis despite different growth conditions. Notably,
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vesicles isolated from strain 168 biofilm were free floating and non-adherent to other material
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whereas strain 3610 biofilm vesicles were often associated to flagella and other debris. This may
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reflect reduced production of exopolysaccharide by strain 168 due to a mutation in the epsC gene
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(McLoon et al., 2011).
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Previously, Tashiro et al showed that OMVs with stronger ζ potentials were more likely
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to associate with cells in P. aeruginosa (Tashiro et al., 2010). B. subtilis cells have a strongly
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negative ζ potential with a magnitude similar to the vesicles recovered from the supernatant.
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This finding is consistent with notion that B. subtilis extracellular vesicles originate from the cell
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membrane and have the potential to interact with bacterial cells (Ahimou et al., 2001).
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When isolating vesicles from the B. subtilis 168 lab and 3610 environmental strains, we
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noted that vesicles isolated from each strain under the same culture conditions resulted in
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different DLS profiles. Strain 168 produced two populations similar in size to vesicles from
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other organisms whereas strain 3610 showed signals not typically associated with vesicles and
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more consistent with the presence of contaminating structures. The apparent paucity of vesicle
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signatures in the DLS profile of vesicle preparation from strain 3610 most likely reflects flagella
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contamination, which could skew the DLS data. For strain 168, vesicles were so abundant
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relative to any contaminating flagella that they exhibited a profile like those reported for other
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vesicle preparations (Rivera et al., 2010). DLS results of sfp+ strains, SL3610 and RL3090, also
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demonstrated similar contaminant distributions to strain 3610 whereas strain AD3610 (sfp-)
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displayed two clean peaks like strain 168. The dimensions obtained from DLS data were
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consistent with measurements from TEM images, which also revealed that more vesicles could
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be recovered from strain 168 compared to strain 3610. Strain 168 vesicle preparations contained
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so many vesicles that they were identified in the entire EM field of view whereas in strain 3610
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vesicle preparations, only an occasional vesicle was present in microscopic fields. It is notable
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that cells from all strains exhibited putative vesicular blebs on the cell membrane in SEM images
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even though we recovered various amounts of vesicles from the strains (data not shown).
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Although such blebs cannot be conclusively identified as vesicles based on sharing similar
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morphology and dimensions, their presence in the surface of wild type strains, and their
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relatively scarcity in the culture supernatants of those strains is consistent with, and supportive of
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the notion that they are made by wild type strains but disrupted by surfactin after release from
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cells.
Accepted Article
329
349
The ability of strain 168 to be easily genetically manipulated as well as to produce large
350
quantities of vesicles makes it an ideal model strain to study the genetics and mechanisms of
351
vesicle production in Gram-positive bacteria. Further, mutations in sfp, epsC, swr,and PdegQ, as
This article is protected by copyright. All rights reserved.
well as the lack of the plasmid-borne gene rapP in strain 168 were identified as genetic
353
differences between strain 168 and strain 3610 (McLoon et al., 2011). We further investigated
354
the effect of the sfp gene on vesiculogenesis. Strain AM373 (168 harboring wild type copies of
355
aforementioned genes) showed decreased amounts of recoverable vesicles compared to the
356
background 168 strain by TEM and metabolic labeling (data not shown). This was evidence that
357
the lack of a functional copy of one or more of these genes could be responsible for high vesicle
358
recovery in the 168 strain. We investigated the candidate gene, sfp, further for its effects on
359
vesicle production in B. subtilis. When a functional copy of sfp was restored into strain 168, the
360
quantity of recoverable vesicles was greatly reduced (RL3090). Furthermore, when sfp was
361
disrupted in strain 3610 (AD3610), vesicle recovery levels increased similar to strain 168. Strain
362
SL3610, a sfp complement strain of AD3610, again showed abrogated vesicle recovery. These
363
results suggest that sfp is an important factor in the amount of vesicles recovered between wild
364
strains and lab strains of B. subtilis and further reinforces the authenticity of vesiculogenesis by
365
the Gram-positive bacterium.
366
recently been reported in mycobacteria (Rath et al., 2013).
Accepted Article
352
We note that genetic regulation of vesicular production has
367
Sfp protein is required for surfactin biosynthesis; therefore strains lacking sfp cannot
368
produce surfactin. Surfactin has been shown to disrupt lipid membranes and so we investigated
369
surfactin as the mechanism behind recovering variable amounts of vesicles between wild and
370
domestic strains of B. subtilis. We were able to recover similar amounts of vesicles from strain
371
RL2663 as from strain 168. RL2663 has a functional sfp gene but a mutation in srfAA, which is,
372
also required for surfactin synthesis. This data led us to believe that surfactin production affected
373
the amount of recoverable vesicles. We confirmed that vesicle incubation in pure surfactin and
374
cell-free supernatant from surfactin-producing strain 3610 leads to less recoverable radioactivity
This article is protected by copyright. All rights reserved.
in the vesicle pellet compared to incubation in surfactin-nonproducing strain 168 supernatant,
376
PBS, BHI broth, and 10% EtOH indicative of vesicle disruption. We also determined that strain
377
3610 supernatant and pure surfactin disrupted vesicles from B. anthracis, suggesting that wild B.
378
subtilis can utilize surfactin to disrupt extracellular vesicles produced by other species. The high
379
stability of B. anthracis vesicles in strain 168 supernatant, which corresponds to depleted BHI
380
media, may be due to the cells utilizing one or more destabilizing molecules in the media but this
381
difference was not pursued further.
382
functions as iron transport, it is conceivable that surfactin-mediated disruption of vesicles from
383
other species provides a competitive advantage to B. subtilis (Rath et al., 2013). In summary,
384
our data implies that environmental strains produce vesicles that are disrupted by surfactin and
385
provides a new function for this lipopeptide in promoting release of vesicular cargo into the
386
extracellular space.
Accepted Article
375
Since vesicles have been implicated in such critical
387
In conclusion, we have characterized extracellular vesicles produced by Bacillus subtilis.
388
These vesicles are not only similar in size and morphology to other bacterial vesicles but we
389
were also able to describe vesiculogenesis in various strains. In addition, the ability to recover a
390
large amount of vesicles from strain 168 makes it an efficient model for the study of
391
vesiculogenesis in Gram-positive bacteria. We have identified a gene, sfp, which is
392
nonfunctional in most laboratory strains, that indirectly affects the quantity of vesicles
393
recoverable from cell cultures. The lipopeptide surfactin, which requires functional sfp, disrupts
394
vesicles in the extracellular milieu to release vesicle cargo. To our knowledge, this is the first
395
time a mechanism for self-and foreign-vesicle cargo release has been reported in Gram-positive
396
bacteria. The occurrence of vesicular production in B. subtilis combined with the ability to
This article is protected by copyright. All rights reserved.
induce release of vesicular content across species boundaries provides a potentially outstanding
398
platform for the dissection of this enigmatic process in a model organism.
399
Experimental Procedures
400
Bacterial Strains
Accepted Article
397
401
Bacillus subtilis 168 strain was obtained from ATCC #23857, and B. subtilis strains
402
NCIB# 3610, AMA373, RL3090, RL2663, and AD3610 were gifts from Richard Losick
403
(Boston, MA) (Table 1). Strain AD3610 was constructed using techniques previously described
404
(Yasbin; Young, 1974; Gryczan et al., 1978; Wach, 1996). Briefly, PCR was utilized to disrupt
405
the sfp gene with a Kan cassette (Table S1). The construct was transformed into strain PY79 and
406
moved into strain 3610 by phage transduction. Strain SL3610 was constructed by ectopic
407
integration of sfp-spec into the AmyE locus of AD3610 by using pDG1730 and In-Fusion HD
408
Cloning Plus kit (Clontech) (Table S1). Transformation was performed as stated above. The lab
409
strain of B. anthracis Sterne 34F2 (pXO1+, pXO2−) used in stability assays was obtained from
410
Alex Hoffmaster at the Center for Disease Control (Atlanta, GA). All strains were grown and
411
maintained from 25% glycerol -80oC frozen stock on brain heart infusion (Difco) agar or broth at
412
37oC. Antibiotics used include 0.5 μg ml-1 erythromycin and 2.5 μg ml-1 lincomycin (mls), 100
413
μg ml-1 spectinomycin, and 10 μg ml-1 kanamycin.
414
Vesicle Isolation
415
Vesicles were obtained with minor modifications of the method used previously for the
416
isolation of B. anthracis vesicles (Rivera et al., 2010). Briefly, bacterial cultures were spun for
417
20 min at 4oC at 15,000 x g to remove cells. The resulting supernatant was then filtered through
418
a 0.22 um Milipore Express Plus Membrane and the filtrate was concentrated using Amicon
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ultrafiltration system (Millipore) with 100 kDa filter. The concentrate was then further filtered
420
through a 0.8 um syringe filter (Corning) to remove larger cellular debris or aggregated material.
421
The filtered supernatant was then centrifuged at 195,000 x g for 1 h at 4oC and washed with
422
phosphate buffered saline (PBS). The vesicle pellet was resuspended in PBS or water. Heat-
423
killed culture vesicles were prepared by autoclaving an overnight culture. The cells were then
424
spun for 20 min at 4oC at 15,000 x g and washed with PBS. The dead cells were then suspended
425
in fresh media and incubated overnight with shaking. Vesicles were then isolated in parallel with
426
live cultures.
427
Vesicle Size by Dynamic Light Scattering (DLS)
Accepted Article
419
428
The size distribution and diameter of vesicles resuspended in PBS or water was measured
429
using 90Plus/BI-MAS Multi Angle Particle Sizing analyzer (Brookhaven Instruments Corp.), as
430
previously described (Eisenman et al., 2009).
431
Transmission Electron Microscopy (TEM)
432
Vesicles were visualized by TEM using negative and positive staining techniques. For
433
negative staining, vesicles were adhered to 300-mesh carbon and Formvar coated Copper grids
434
for 2 min, stained with 1% phosphotungstic acid and viewed on a JEOL 100CXII or 1200EX at
435
80kV.
436
paraformaldehyde in 0.1 M sodium cacodylate buffer. Samples were then postfixed with 1%
437
osmium tetroxide followed by 2% uranyl acetate en-bloc staining. The samples were dehydrated
438
through a graded series of ethanol and embedded in LX112 resin (LADD Research Industries,
439
Burlington VT). Pellicle biofilms were fixed for 24 h with 2.5% gluteraldehyde, 0.2 M sodium
440
cacodylate, 0.4 M sucrose, and 10mM MgCl2 (pH 7.4) in a 1:1 ratio with the MSgg culture media.
For embedded TEM, vesicles were fixed with 2.5% glutaraldehyde and 2%
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Pieces of each fixed biofilm were processed in suspension on a rotator. The biofilm samples were
442
postfixed with 1% osmium tetroxide, 0.7% potassium ferrocyanide for 1 h followed by 1% uranyl
443
acetate. The biofilm samples were dehydrated through a graded series of ethanol and then
444
embedded in LX112 resin (LADD Research Industries, Burlington VT). Ultrathin (80 nm) sections
445
of all samples were cut on a Reichert Ultracut UCT, stained with uranyl acetate followed by lead
446
citrate.
447
Scanning Electron Microscopy (SEM)
Accepted Article
441
448
Cells were fixed in 5% glutaraldehyde, 0.2 M sodium Cacodylate, 0.4 M Sucrose, 10mM
449
MgCl2 pH 7.4 and mixed 1:1 with suspension. The biofilm samples were postfixed for 30 min with
450
1% osmium tetroxide, 0.7% potassium ferrocyanide, 0.1 M sodium cacodylate, 0.2 M sucrose, and
451
5 mM MgCl2 (pH 7.4). Samples were dehydrated through a graded series of ethanol and critical
452
point dried using liquid carbon dioxide in a Tousimis Samdri 795 Critical Point Drier (Rockville,
453
MD). The samples were sputter coated with gold-palladium in a Denton Vacuum Desk-2 Sputter
454
Coater (Cherry Hill, NJ). Prepared cells were then examined in a Zeiss Supra Field Emission
455
Scanning Electron Microscope (Carl Zeiss Microscopy, LLC North America), using an accelerating
456
voltage of 5 KV.
457
Glucuronoxylomannan Immunogold TEM
458
The polysaccharide, glucuronoxylomannan (GXM), was isolated from Cryptococcus
459
neoformans and fractionated to less than 10 kDa in size by filtration through Amicon Ultra
460
Centrifugal Filters (Millipore). GXM was added to a culture of strain 168 at inoculation and
461
incubated at 37oC for 18 h. Vesicles were then isolated and either sonicated (20 sec sonication
462
and 30 sec incubation on ice, repeated 4 times) or not (Sonic Dismembrator Model 100; Fisher
This article is protected by copyright. All rights reserved.
Scientific). Vesicles were fixed with 2% paraformaldehyde, 0.1 % glutaraldehyde in 0.1 M
464
Phosphate buffer, followed by dehydration with a graded series of ethanol, and a progressive
465
lowering of the temperature to -20° in a RMC FS 7500. Samples were then embedded in
466
Lowicryl HM-20 monostep resin (Electron Microscopy Sciences) and polymerized using UV
467
light. Ultrathin sections were cut on a Reichert Ultracut UCT.
Accepted Article
463
468
Immunogold labeling was performed by fixing the samples in 4% paraformaldehyde
469
(Electron Microscopy Sciences), 0.1% glutaraldehyde (Polysciences), and buffered in 0.1 M sodium
470
cacodylate (pH 7.2). Samples were incubated with the primary monoclonal antibody (mAb) 18B7
471
against GXM diluted in incubation buffer overnight at 4oC. The sample was washed with BSA-C
472
(Electron Microscopy Sciences) and incubated with 10 nm immunogold-conjugated secondary
473
antibody (donkey anti-mouse) (Electron Microscopy Sciences) for 2 h at room temperature.
474
Samples were washed again and postfixed with 2% gluteraldehyde/PBS. Finally, samples were
475
stained with 4% uranyl acetate and imaged using JEOL 100CXII transmission electron microscope
476
at 80kv. Controls included preparations with no primary antibody as well as preparations with an
477
irrelevant IgG antibody.
478
Metabolic labeling with C-14 Palmitic Acid
479
B. subtilis and B. anthracis cells were labeled with radioactivity as previously described
480
for C. neoformans with minor modifications (Wolf et al., 2012). Briefly, an overnight culture of
481
bacteria was diluted to a suspension with an optical density at 600 nm (OD600) of ~0.1. 1-2 μCi
482
of 14C PA was added to aliquots of 15-30 ml. The samples were incubated at 37oC until OD600
483
doubled to ~0.2 and the cells were then pelleted, washed, and resuspended in fresh BHI.
484
Aliquots of 1 ml were added to 14 ml BHI in PETG flasks (Thermo Scientific) and grown
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overnight at 37oC. Overnight cultures were normalized by OD600 and vesicles were harvested by
486
centrifuging cells for 20 min at 15,000 x g at 4 oC and filtering the supernatant with a 0.8 μm
487
syringe filter (Corning) and the resultant supernatant was concentrated with Amicon Ultra
488
Centrifugal Filters 100 kDa (Millipore). The concentrate was spun at 195,000 x g for 1 h at 4 oC,
489
and washed with PBS. The vesicle pellet was then resuspended in PBS for dynamic light
490
scattering and scintillation counting (CPM). Controls included B. subtilis incubated without 14C
491
PA, cells incubated with 150 mM NaN3, and cultures without B. subtilis cells.
492
production over time was studied by counting radioactivity of vesicles isolated at 0, 4, 12, and 18
493
h. Samples measured include cell pellets, supernatant, concentrate FT, concentrate supernatant,
494
and vesicle pellet and each fraction represented as a ratio of the total sample. Scintillation
495
counting was done in a Packard Bioscience Tricarb A2900 liquid scintillation counter. The
496
position of vesicle fractions was determined by resuspending labeled vesicles in 30% Optiprep
497
Density Gradient Medium (Sigma) and layering 2 ml of 25, 20, 15, and 10% Optiprep above the
498
vesicle sample. The gradient was spun for 16 h at 140,000 x g at 4oC. 0.5 ml fractions were
499
taken from the top of the gradient and counted.
500
Vesicle Stability
Accepted Article
485
501
Vesicle stability over time, incubated in PBS, was measured by incubating
Vesicle
14
C PA-
502
labeled vesicle pellets at room temperature for 0, 24, 48 and 72 h in PBS followed by
503
ultracentrifugation and counting of supernatants and pellets. Vesicle stability was measured in
504
10 μg ml-1 and 100 μg ml-1 surfactin (resuspended in EtOH at 10 mg ml-1) (Sigma), BHI media,
505
PBS, 10% EtOH, and cell-free supernatants from B. subtilis strains 168 and 3610 week old
506
cultures. B. anthracis stability was measured similarly to B. subtilis but with the added condition
507
of cell-free supernatant from B. anthracis strain Sterne 34F2. Vesicle stability was measured by
This article is protected by copyright. All rights reserved.
14
C PA-labeled vesicle pellets resuspended in the various conditions at 37 oC on a
incubating
509
nutator and after 24 h, samples were ultracentrifuged and radioactivity of supernatants and
510
pellets were counted.
511
ζ Potential Measurements
Accepted Article
508
512
The ζ potential of vesicles in water with 10mM dextrose and 1mM KCl was calculated
513
using a ζ potential analyzer (ZetaPlus; Brookhaven Instruments Corp.).
514
Protein Characterization
515
Vesicles were isolated and purified by density gradient centrifugation using OptiPrep and
516
vesicle containing fractions were dialyzed overnight against PBS at 4oC with one buffer change.
517
Cells were grown for 18 h at 37o, spun down and washed with PBS. Cell membrane fractions
518
were isolated as previously described (Petrackova et al., 2010).
519
lyophilized and analyzed.
All samples were then
520
Proteins were precipitated from vesicle suspensions, membrane fractions, and bacterial
521
lysates by adding six volumes of ice-cold acetone. After incubation overnight, the acetone was
522
removed and the pellets dissolved in 25 mM ammonium bicarbonate. Proteins were reduced with
523
10 mM DTT at room temperature for 1 h and the cysteine residues were subsequently alkylated
524
with 10 mM iodoacetamide at room temperature for 1h in the dark. Protein digestion was carried
525
out with trypsin (Promega, 1:20, w/w) at 37ºC overnight. The peptide mixtures from in-solution
526
tryptic digestions were analyzed using nanoflow LC-MS/MS. The peptides were loaded onto a
527
0.3 x 5 mm C18 precolumn (New Objective) and then eluted with a linear gradient of 2-90%
528
acetonitrile in 0.1% aqueous solution of formic acid. The gradient was delivered over 120 min by
529
a nanoLC-1Dplus (Eksigent) at a flow-rate of 200 nl/min, through a 75-µm x 15 cm fused silica
This article is protected by copyright. All rights reserved.
capillary C18 HPLC PepMap column (LC Packings) to a stainless steel nano-bore emitter
531
(Proxeon). The peptides were scanned and fragmented with an LTQ XL linear ion trap mass
532
spectrometer (ThermoFinnigan) operated in data-dependent and MS/MS switching mode using
533
the three most intense precursors detected in a survey scan from 400 to 1600 amu (three µscans).
534
Normalized collision energy was set to 35%, and dynamic exclusion was applied during 3 min
535
periods to avoid repetitive fragmentation of ions. Generated .raw files were converted to .dta
536
files for Mascot database. A database containing the NCBI B. subtilis (4176 sequences, as of
537
January 20, 2012) was searched using Mascot Software (version 2.3, Matrix Science) for protein
538
identification. Search criteria included trypsin specificity with one missed cleavage allowed, and
539
methionine oxidation as variable modification. A minimum precursor and fragment-ion mass
540
accuracy of 1.2 and 0.3 Da, respectively, and a requirement of at least one bold red peptide (i.e.
541
highest scoring peptide matching to protein with highest total score). Cut-off values for Mascot
542
scores of peptides and proteins were set to 45 (p < 0.05) and 51 (p < 0.01), respectively, for
543
considering them as being accurate identifications. Proteins identified with only one peptide
544
were inspected manually. The false positive rate (1,23%) was calculated searching the same
545
spectra against the NCBI B. subtilis decoy database, using the version updated January 20, 2012.
546
A combined list of proteins identified in all replicates was condensed in order to remove
547
redundant IDs such as orthologous sequences, redundant database entries, and indistinguishable
548
isoforms based on observed peptide coverage.
Accepted Article
530
549
Vesicle preparation protein content from strains 168, 3610, and RL2663 was normalized
550
with Pierce BCA Protein Assay Kit (Thermo Scientific) and 7.5 ug of protein was run on 4–20%
551
Mini-PROTEAN® TGX Stain-Free Gel (Bio-Rad) and silver stained using Pierce Silver Stain
552
Kit (Thermo Scientific).
This article is protected by copyright. All rights reserved.
Biofilm Assay and Vesicle Isolation
Accepted Article
553 554
Biofilms were grown as described in McLoon et al. with modifications (McLoon et al.,
555
2011). Bacteria were grown from frozen stock on BHI agar overnight at 37oC. A single colony
556
was then inoculated into BHI broth for 4 h at 37oC with shaking. The culture was inoculated
557
(1:1000) into MSgg media (100 mM MOPS, pH 7, 5 mM potassium phosphate, pH 7, 700 μM
558
CaCl2, 2 mM MgCl2, 50 μM FeCl3, 50 μM MnCl2, 1 μM ZnCl2, 2 μM thiamine, 50 μg ml-1
559
tryptophan, 50 μg ml-1 phenylalanine, and 50 μg ml-1 threonine, 0.5% glycerol and 0.5%
560
glutamate). Biofilms were grown at 30oC for either 4 or 8 d without shaking. After the desired
561
time, the biofilm culture was disrupted and centrifuged at 15,000 x g for 15 min to remove
562
cellular debris. The resulting supernatant was filtered through a 0.22 um Milipore Express Plus
563
Membrane and then concentrated using Amicon Ultra Centrifugal Filters 100 kDa (Millipore).
564
The resultant concentrate was centrifuged at 195,000 x g at 4 oC and washed with PBS. The
565
vesicle pellet was resuspended in PBS or water. Whole biofilm pellicles were submitted to TEM
566
and SEM preparations.
567
Statistics
568
Statistics were performed in Prism (GraphPad Software). Error bars indicate mean and
569
standard deviation. Comparisons between ratio of natural and sonicated vesicles containing gold
570
particles was done using Chi2 test.
571
compared using an unpaired t-test. One-way ANOVA was performed comparing vesicle density
572
and diameter. Vesicle pellet radioactivity between strains 168 and 3610 was analyzed with an
573
unpaired t-test. A one-way ANOVA was utilized when comparing vesicle pellet radioactivity
574
among strains. One-way ANOVA was also used to compare stability of pellet radioactivity after
The diameters of natural and sonicated vesicles were
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incubation in various conditions. Significance was indicated as follows: p
Accepted Article
Extracellular Vesicles Produced by the Gram-positive Bacterium Bacillus subtilis are Disrupted
2
by the Lipopeptide Surfactin
1
3 4
Running Title: Vesicle Production and Disruption in Bacillus subtilis
5 6
Lisa Brown1*, Anne Kessler1, Pablo Cabezas-Sanchez3, Jose L. Luque-Garcia3, Arturo
7
Casadevall1,2
8 9
1
Department of Microbiology and Immunology, Albert Einstein College of Medicine, Bronx, NY
10
10461, USA
11
2
Department of Medicine, Albert Einstein College of Medicine, Bronx, NY 10461, USA
12
3
Department of Analytical Chemistry, Universidad Complutense de Madrid, Madrid, Spain.
13 14
*For correspondence E-mail: [email protected]; Tel: (+1) 718 430 3730; Fax:
15
(+1) 718 430 8701.
16 17 18
This article has been accepted for publication and undergone full peer review but has not been through the copyediting, typesetting, pagination and proofreading process, which may lead to differences between this version and the Version of Record. Please cite this article as doi: 10.1111/mmi.12650 This article is protected by copyright. All rights reserved.
Accepted Article
19 20
Summary/Abstract
21
Previously, extracellular vesicle production in Gram-positive bacteria was dismissed due
22
to the absence of an outer membrane, where Gram-negative vesicles originate, and the difficulty
23
in envisioning how such a process could occur through the cell wall. However, recent work has
24
shown that Gram-positive bacteria produce extracellular vesicles and that the vesicles are
25
biologically active. In this study, we show that Bacillus subtilis produces extracellular vesicles
26
similar in size and morphology to other bacteria, characterized vesicles using a variety of
27
techniques, provide evidence that these vesicles are actively produced by cells, show differences
28
in vesicle production between strains, and identified a mechanism for such differences based on
29
vesicle disruption. We found that in wild strains of B. subtilis, surfactin disrupted vesicles while
30
in laboratory strains harboring a mutation in the gene sfp, vesicles accumulated in the culture
31
supernatant.
32
anthracis, indicating a mechanism that crossed species boundaries. To our knowledge, this is the
33
first time a gene and a mechanism has been identified in the active disruption of extracellular
34
vesicles and subsequent release of vesicular cargo in Gram-positive bacteria. We also identify a
35
new mechanism of action for surfactin.
36
Introduction
Surfactin not only lysed B. subtilis vesicles, but also vesicles from Bacillus
37
In recent years, several laboratories have reported that bacteria, fungi, and parasites
38
produced extracellular vesicles, which in some instances contain proteins associated with
39
virulence.
40
focused on the outer membrane vesicles (OMV) produced by Gram-negative bacteria (Kuehn;
The most extensive studies of extracellular vesicle production in bacteria have
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Kesty, 2005; Ellis; Kuehn, 2010; Kulp; Kuehn, 2010).
42
concentrated and protected manner, and therefore are potent virulence factors of pathogenic
43
Gram-negative bacteria. These vesicles contain toxins, DNA, immunomodulatory compounds,
44
communication factors, and adhesins and have been associated with cytotoxicity, bacterial
45
attachment, intercellular DNA transfer, and invasion (Dorward et al., 1989; Kuehn; Kesty, 2005;
46
Ellis; Kuehn, 2010; Maldonado et al., 2011). Until recently, vesicle production by Gram-
47
positive bacteria was overlooked due to the inference that differences in the cell wall structure of
48
Gram-negative and Gram-positive bacteria precluded their release.
49
bacteria, the cell wall of Gram-positive bacteria consists of primarily peptidoglycan and does not
50
have an outer membrane. However, the discovery of extracellular vesicles in fungi, which also
51
have large cell walls, catalyzed the search of vesicular structures in Gram-positive bacteria
52
(Rodrigues et al., 2007).
Accepted Article
41
OMVs can release materials in a
Unlike Gram-negative
53
The first evidence for vesicles in Gram-positive bacterial cells was provided by Dorward
54
and Garon but they were unable to establish function (Dorward; Garon, 1990). More recently,
55
vesicle production was described in Mycobacterium ulcerans, Staphylococcus aureus,
56
Mycobacterium tuberculosis, and Bacillus anthracis (Marsollier et al., 2007; Lee et al., 2009;
57
Rivera et al., 2010; Gurung et al., 2011; Hong et al., 2011; Prados-Rosales et al., 2011; Thay et
58
al., 2013). M. tuberculosis vesicles contain compounds that modulate TLR2 responses while
59
those of B. anthracis contain anthrax toxin components, indicating that Gram-positive membrane
60
vesicles are potentially involved in pathogenesis (Rivera et al., 2010; Prados-Rosales et al.,
61
2011). This observation suggested that Gram-positive cells, like Gram-negative bacteria, utilized
62
vesicle secretion to deliver concentrated amounts of toxins and other compounds into the
63
extracellular space (Marsollier et al., 2007; Lee et al., 2009; Rivera et al., 2010; Prados-Rosales
This article is protected by copyright. All rights reserved.
et al., 2011; Hong et al., 2011; Gurung et al., 2011; Thay et al., 2013). The disruption of B.
65
anthracis vesicles by serum was identified as a potential mechanism for vesicle cargo release, as
66
it is important to understand what happens to vesicles and their contents after release from the
67
cell (Wolf et al., 2012). Recently, vesicle production has also been described for Streptococcus
68
pneumonia and pneumococcal vesicles are used to release pneumolysin (Olaya-Abril et al.,
69
2014).
Accepted Article
64
70
In this work, we explored vesicle production in the Gram-positive, endospore-forming
71
bacterium, Bacillus subtilis. B. subtilis is considered the model system for Gram-positive
72
bacteria and is therefore an excellent organism to study the genetic and cellular machinery
73
involved in Gram-positive vesiculogenesis. Two common strains of B. subtilis studied were the
74
3610 environmental strain, which forms robust biofilms and the laboratory strain 168, which has
75
attenuated biofilm production. Both strains produced varying amounts of recoverable vesicles
76
under different conditions. Environmental strains of B. subtilis are often not easily genetically
77
manipulated; therefore domesticated lab strain 168 was created to be easily transformable and
78
used for genetic studies in Gram-positive bacteria (Earl et al., 2007). It is hypothesized that both
79
strains 168 and 3610 are descendants of B. subtilis Marburg strain and vary from each other by
80
only 22 SNPs (Zeigler et al.,2008). These limited genetic differences and ability to isolate
81
differing amounts of vesicles make these two strains ideal to study the mechanism of vesicle
82
production in B. subtilis. McLoon et al identified five of these genetic differences between
83
strains 168 and 3610 that are involved in biofilm formation (mutations in 168 in the genes sfp,
84
epsC, swrA, PdegQ, and lack of the plasmid-borne gene rapP) (McLoon et al., 2011). Further
85
study of these two strains has led us to identify the gene, sfp, which affects the amount of
86
vesicles recovered in B. subtilis. Lab strains, including strain 168, harbor a mutation in sfp
This article is protected by copyright. All rights reserved.
resulting in a truncated, nonfunctional protein, whereas environmental strains, like strain 3610,
88
harbor a wild type copy of this gene (Nakano et al., 1988).
Accepted Article
87
89
Sfp is a 4’-phosphopantetheinyl transferase which transfers a phosphopantetheine group
90
from coenzyme A to a peptidyl carrier protein during biosynthesis of the lipopeptide, surfactin,
91
and the siderophore, bacillibactin, in B. subtilis (Nakano et al., 1988; Quadri et al., 1998; May et
92
al., 2001). We found that strains harboring a functional sfp gene produced very few recoverable
93
vesicles whereas strains with nonfunctional copies of sfp, produced a massive amount of
94
recoverable vesicles. To determine the mechanism behind this difference in vesicle quantity, and
95
how sfp was affecting it, we explored the effect of surfactin on vesicles. We found that vesicles
96
from B. subtilis and B. anthracis were indeed disrupted by the presence of surfactin. And this
97
was not surprising considering surfactin can perturb lipid membranes (Bernheimer; Avigad,
98
1970; Heerklotz; Seelig, 2001; Carrillo et al., 2003). This data suggests that the production of
99
surfactin serves as a mechanism for vesicular cargo release in B. subtilis, as well as the ability to
100
disrupt vesicles from B. anthracis, indicating a mechanism that reaches across species barriers.
101
This work sets the stage for dissecting the function of vesiculogenesis in a genetically tractable
102
Gram-positive bacterium.
103
Results
104
Bacillus subtilis Produces Extracellular Vesicles
105
We recovered vesicles from the culture supernatants of Bacillus subtilis lab strain 168,
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wild type strain 3610, AM373, RL3090, AD3610, SL3610, and RL2663 (Table 1).
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morphology and color of the vesicle pellet from each strain was the first indication that there
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were differences in vesicle preparations among strains. Centrifugation of strain 168 culture
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The
supernatants produced a large, reddish-brown vesicle pellet whereas centrifugation of strain 3610
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culture supernatant and 168 heat-killed preparations produced only small, clear pellets (Fig. 1A-
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C). A massive amount of vesicles from strain 168 were visualized by both negative and positive
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staining transmission electron microscopy (TEM) (Figure 1D-E). In contrast, very few vesicles
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were identified in electron micrographs of identical preparations of vesicles from strain 3610.
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Due to the abundance of vesicles in the supernatant of strain 168, most of the subsequent
115
characterization was performed on vesicles isolated from this strain.
Accepted Article
109
116
Given the propensity of lipids to organize into vesicle-like structures, considerable effort
117
was devoted to establishing that recovered vesicles were not artifacts of broken cells or a result
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of self-assembly by extracellular lipids. Consistent with a requirement for cell viability in vesicle
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production, we observed very few vesicles by negative staining TEM when heat-killed 168 cells
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were added to bacterial media, allowed to incubate at 37o C for 18 h in a shaker, and then
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processed for vesicle preparation under the same protocol (data not shown).
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To exclude the possibility that the recovered vesicles were the result of spontaneous
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extracellular assembly of secreted lipids we performed an experiment where the fungal
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polysaccharide glucuronoxylomannan (GXM), of Cryptococcus neoformans, was added to the
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bacterial culture and then its presence and absence inside vesicles was measured by immunogold.
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To ensure that the GXM was small enough in size to become encapsulated inside vesicles it was
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fractionated to less than 10 kDa in size by ultrafiltration through selective molecular mass
128
membranes. This size retains reactivity with specific monoclonal antibodies (mAbs). We
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reasoned that if vesicles were forming spontaneously from lipids, they would contain far more
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GXM than vesicles assembled by the bacterial cell (for a scheme of the experimental design see
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Figure 2A), and that was precisely what was measured. As a control, we sonicated vesicles
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isolated from cultures incubated with GXM, expecting that these would contain more GXM,
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since subsequent self-assembly in the presence of GXM would result in trapping of the
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polysaccharide in newly formed vesicles. TEM images of the natural vesicle preparations
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showed the presence of GXM in the sample (Figure 2B).
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significantly more GXM was inside vesicles that reformed after sonication compared to natural
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vesicles produced from live bacterial cultures (6.9% and 2.9%, respectively) (Figure 2C). These
138
results provided strong support for the notion that vesicles are produced by the cell and are not
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the result of spontaneous organization of lipids in the media. Sonicated vesicle diameter was also
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significantly smaller and varied less than that of natural vesicles, indicating another difference
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between B. subtilis-produced vesicles and those that reformed from the aggregation of lipids
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(94.54 ± 1.869 and 115.4 ± 2.69) (Figure 2D). We conclude that B. subtilis, like other Gram-
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positive bacteria, actively produce extracellular vesicles (Marsollier et al., 2007; Lee et al., 2009;
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Rivera et al., 2010; Prados-Rosales et al., 2011).
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Physical Characterization of B. subtilis Vesicles
Accepted Article
132
In accordance to predictions,
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Vesicle preparations from planktonic B. subtilis cultures were studied using dynamic
147
light scattering (DLS) to determine the approximate size of the vesicles. DLS revealed two
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populations of approximately 50 and 150-250 nm in preparations from strain 168 (Figure 3B).
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This size distribution is similar to that reported for vesicles produced by other bacteria (Kuehn;
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Kesty, 2005). DLS analysis of vesicle preparations from B. subtilis strain 3610 under the same
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conditions revealed less distinct distributions and the presence of larger structures, which may be
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flagella given their abundance in TEM micrographs.
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Analysis of strain 168 supernatant pellets by TEM with negative staining revealed large
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numbers of vesicles (Figure 1D) whereas very few vesicles were recovered from strain 3610
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when isolated from the same culture conditions. Images of strain 168 vesicles obtained by
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embedded TEM were measured using ImageJ software, and the result was a mean diameter of
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137.7 nm (Figure 1E). However, because vesicles are spherical, and TEM represents transverse
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sections, a 1.27 correction factor for these 2D measurements was utilized as suggested by Kong
159
et al to estimate the actual equatorial dimensions (Kong et al., 2005). With the 1.27 correction
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factor, the mean diameter of vesicles was 174.4 nm (Figure 3A). It is noteworthy that when DLS
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and TEM measurements are compared, TEM tends to underestimate size while DLS
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overestimates the size of samples (Egelhaaf et al., 1996; Hong et al., 2011). TEM images of
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strain 168 vesicles indicated that some vesicles were more electron dense than others. Smaller
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vesicles were significantly more electron dense than larger vesicles (Figure 3C).
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diameters were categorized visually in three groups based on electron density with mean
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diameters of 214.5 nm for least dense (+), 152.2 nm for medium dense (++), and 120.4 nm for
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most dense (+++) (Fig. 3D-F).
Accepted Article
153
Vesicle
168
Spherical protrusions from bacterial cells with dimensions similar to vesicles were
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observed in electron micrographs of B. subtilis cells. TEM images of strain 168 cells showed
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rounded structures strongly suggestive for putative vesiculation in the cell membrane (Figure
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4A-B).
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staining TEM in various stages of release (Figure 4C). Finally, SEM images of strain 168 cells
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also showed multiple protrusions on cells consistent with the emergence of vesicles (Figure 4D).
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Although we recovered various quantities of vesicles from strains 168 and 3610, it is notable that
Structures similar to vesicles were also identified on strain 168 cells by negative
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the number of putative vesicle protrusions on cells was not significantly different (data not
176
shown).
Accepted Article
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177
Bacterial cells were labeled with
14
C palmitic acid (14C PA), which is incorporated into
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vesicles, to study stability, production over time, and to compare recoverable vesicle quantity
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among strains (Rivera et al., 2010; Prados-Rosales et al., 2011; Wolf et al., 2012). B. subtilis
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strain 168 vesicles were isolated after 0, 4, 12, 18, and 24 h of growth and radioactivity of the
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vesicle pellets were analyzed as fractions of the total culture. Strain 168 manifested increased
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vesicle production over time, which leveled off after 12 h of growth (Figure 5A). Stability of
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strain 168 vesicles in PBS was also measured by
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incubated for 0, 24, 48 and 72 h before being spun down, counted and the ratio of vesicle versus
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supernatant radioactivity was analyzed. Vesicle pellets contained high levels of radioactivity for
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24 h but subsequently decreased, indicating that vesicles were stable for approximately 24 h
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(Figure 5B). DLS measurements of vesicle stability preparations from each time point showed
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consistent peaks of approximately 50 and 100-150 nm in diameter (Figure 5C-F).
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Chemical Composition of Vesicles
14
C PA labeling. Vesicle preparations were
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Proteomic analysis was performed on strain 168 vesicles, cell membrane fractions, and
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cells. B. subtilis 168 vesicles were enriched in 30 proteins and shared 46 with both the cell and
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cell membrane fractions (Figure 6A). Vesicles were enriched in lipoproteins and contained many
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siderophore-binding proteins (Table S2). SunI, the sublancin immunity protein, was found
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associated with the vesicles (Dubois et al., 2009). Most of the proteins in the vesicle preparation
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were membrane proteins, suggesting that the vesicles originated from the membrane of the
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bacterial cell (Table S2). SDS-PAGE gel of vesicle preparations from strains 168, 3610, and
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RL2663 exhibited similar profiles with only a few differing bands (Figure 6B).
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Vesicle Membrane Potential
Accepted Article
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The ζ potential of strain 168 vesicle preparations were determined to be -54.2917 +/- 9.24
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and -36.724 +/- 10.2 in 10 mM Dextrose and 1 mM KCl, respectively (Figure 6C).
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Vesicles are Constituents of Biofilms
202
Negative staining TEM revealed the presence of vesicle-like structures in B. subtilis
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biofilms. After 4 and 8 d of growth, vesicles were isolated from disrupted B. subtilis strains 168
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(i-ii) and 3610 (iii-iv) biofilm pellicles. Despite the previously mentioned differences in both
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recovered vesicle quantity and biofilm growth between the two strains, similar amounts of
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vesicles were isolated from all cultures and visualized using electron microscopy with negative
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staining (Figure 7A). The size distribution of the vesicles isolated from strains 168 and 3610
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biofilms was determined by DLS, which revealed the presence of two distinct size populations
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across all biofilm vesicle samples with measurements comparable to those described for
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supernatant vesicles from planktonic cultures (Figure 7B).
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Samples of intact B. subtilis strains 168 and 3610 biofilm were further visualized using
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TEM and SEM. Embedded biofilm TEM images of strain 3610 biofilm showed vesicles as part
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of the biofilm matrix (Figure 7C). For SEM, biofilm samples were prepared with ferrocyanide in
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a process that ultimately peels back the outer layer of the biofilm to allow for viewing of the
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internal biofilm constituents. SEM images generated from the ferrocyanide-treated samples
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indicated the presence of putative vesicular events on cells within the B. subtilis strain 3610
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biofilm (Figure 7D). This article is protected by copyright. All rights reserved.
Vesicle Quantification
Accepted Article
218 219
A dramatic difference in vesicle quantity between strains 168 and 3610 was apparent in
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TEM micrographs. To further explore this difference, we utilized metabolic labeling to gain
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more quantifiable information about vesicles from various strains. Cells from strains 168 and
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3610 were labeled with 14C PA and then we measured radioactive counts in cells, flow through
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(FT), supernatant, and vesicle pellets, with each measurement determined relative to the total
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sample (Figure 8A-B). Strain 168 had significantly more radioactivity incorporated into the
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vesicle pellet than strain 3610, even though both strains took up a similar amount of radioactivity
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in the initial labeling step (Figure 8C). This data, along with the DLS and electron micrographs,
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provides further support for the observation that the laboratory strain 168 produces more
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recoverable vesicles than environmental strain 3610.
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Metabolic labeling and density gradient centrifugation was utilized to compare vesicle
230
quantity and density among various B. subtilis strains (Table 1). Cultures of each strain were
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pulsed with
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labeled samples were also recovered and counted; all fraction radioactivities are represented as a
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percentage of the whole sample.
14
C PA and vesicles were isolated. Cells, FT, and supernatant fractions of the
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Strains 168 and AD3610 showed a significantly higher percentage of radioactivity levels
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in the vesicle pellet than pellets of strains 3610, SL3610, and RL3090 (Figure 8D). This indicates
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that supernatants from strains 168 and AD3610 contain a larger quantity of recoverable vesicles
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than 3610, SL3610, and RL3090. Strains 168 and AD3610 have a nonfunctional sfp gene,
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whereas strains 3610, SL3610, and RL3090 harbor fully functional copies of the gene.
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The vesicle preparations were subjected to an Optiprep density gradient and subsequently
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0.5 ml fractions were recovered (Figure 8E). Fractions containing the majority of radioactivity
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were 6, 7, and 8, and the presence of vesicles in these fractions was confirmed by TEM. Outer
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membrane vesicles from Pseudomonas aeruginosa were also found in similar fractions (Tashiro
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et al., 2010).
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Disruption of Vesicles by Surfactin
Accepted Article
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245
We found that we were able to recover large amounts of vesicles from strains that were
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unable to produce surfactin due to a mutation in sfp and very few vesicles from strains with a
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functional sfp gene. This association suggested to us the possibility that vesicle quantities and
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surfactin expression were linked. The gene, sfp, is involved in the biosynthesis of the lipopeptide
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antibiotic, surfactin, in B. subtilis. Because surfactin is known to disrupt membranes, we sought
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to determine if the presence of surfactin in these sfp+ strains resulted in lower amounts of
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recoverable vesicles. Strain RL2663 (3610 background) has a functional sfp gene but a mutation
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in srfAA, an important module of surfactin biosynthesis, rendering the strain unable to produce
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surfactin (Kearns; Losick, 2003). We were able to recover similar amounts of vesicle pellet
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radioactivity from strain RL2663 as strain 168 (Figure 8F), showing the same phenotype for two
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mutations in the surfactin pathway. We also confirmed the large amount of vesicles by TEM
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(data not shown). This result suggests that the presence of surfactin disrupts vesicles, leading to
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less recoverable reactivity in the vesicle pellet fraction.
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To ascertain whether surfactin action was indeed the mechanism behind the differences in 14
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recoverable vesicles among strains, we incubated
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168 in BHI media, strain 168 cell-free supernatant, strain 3610 cell-free supernatant, PBS, 10%
C PA–labeled vesicles isolated from strain
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EtOH, 10 μg ml-1 surfactin, and 100 μg ml-1 surfactin overnight at 37o C on a nutator and then
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recovered the vesicle pellet and spent supernatants.
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radioactivity was significantly reduced when incubated in supernatant from strain 3610 (surfactin
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competent) but stable in strain 168 supernatant (surfactin deficient) (Figure 9A). We also
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recovered less radioactivity in the pellets of vesicles incubated with purified surfactin and the
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data showed a dose response (Figure 9B). These results indicate that the presence of surfactin
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de-stabilizes vesicles and is responsible for the inability to recover large amounts of vesicles
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from surfactin-producing strains of B. subtilis.
Accepted Article
261
We found that recoverable vesicle
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To explore whether the mechanism of vesicle disruption by surfactin extended across
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bacterial species we studied the effect of surfactin on vesicles isolated from B. anthracis. We
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found that pellet radioactivity of
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after incubation with strain 3610 supernatant and purified surfactin (Figure 9C-D). It is notable
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that vesicles from B. anthracis are much less stable under these conditions when compared to B.
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subtilis. B. anthracis vesicle instability was also reported previously (Wolf et al., 2012). We also
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performed the reverse experiments, incubating strain 168 in week-old B. anthracis Sterne,
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Pasteur, and DeltaT cell-free supernatants and did not see a significant change in pellet
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radioactivity compared to BHI and strain 168 supernatant controls (data not shown). This
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indicates B. anthracis does not actively produce a vesicle destabilizing molecule. This result
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extends the role of surfactin to disruption of vesicles from other bacterial species.
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Discussion
281
14
C PA–labeled B. anthracis vesicles significantly decreased
The production of extracellular vesicles appears to be a common phenomenon in both
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Gram-positive and Gram-negative bacteria.
283
vesicles, the quantity of vesicles varies with the strain, that the gene, sfp, influences vesicle
Here, we established that B. subtilis produces
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recovery through the effect of its product surfactin, and surfactin disrupts vesicles produced by
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different species, suggesting a role for unloading vesicles. These results establish B. subtilis as a
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system for studying the physiology of Gram-positive vesicular production.
Accepted Article
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287
Any new description of vesiculogenesis by microorganisms must rule out the possibility
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that such vesicles are artifacts of lipid self-assembly and/or damaged membranes resealing into
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spherical structures. Consequently, we devoted considerable effort to establishing that natural
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vesicles were indeed produced from living cells.
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membrane derived vesicles from several lines of evidence: 1) vesicles were recovered from live
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and not dead cells; 2) metabolic labeling with
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radioactivity into vesicle preparations; 3) the vesicles described for B. subtilis had similar
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dimensions to those produced by other microorganisms; 4) vesicle preparations manifested
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similarities and differences in protein content to membranes consistent with both a common
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origin and a separate synthetic pathway; 5) electron microscopy revealed protrusions from cells
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with dimensions comparable to those of extracellular vesicles suggesting that both are the same;
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6) differences in vesicle recovery among B. subtilis strains implied strain-related and -regulated
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differences in vesicle fate; 7) significantly reduced incorporation of GXM in naturally produced
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vesicles (Rodrigues et al.,2007; Marsollier et al.,2007; Lee et al.,2009; Rivera et al.,2010;
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Prados-Rosales et al.,2011; Maldonado et al.,2011) . The latter experiment was designed to
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show that these vesicles were not forming spontaneously in media, which was assessed by taking
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advantage of a fungal polysaccharide for which mAbs suitable for immunogold labeling of
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vesicle preparations are available. We predicted that if vesicles were forming spontaneously by
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the self-assembly of free lipids that we would find a similarly large amount of vesicles
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containing gold particles in both natural and sonicated vesicles. This, however, was not the case,
14
We conclude that B. subtilis produces
C PA
resulted in the incorporation of
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as sonicated vesicles contained significantly more GXM encapsulated within the vesicles.
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Additionally, these structures were smaller and less variable in size than natural vesicles
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recovered from bacterial supernatants. Furthermore, we noted diameter differences between
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sonicated vesicles that presumably had the opportunity to reassemble and naturally recovered
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vesicles, providing further evidence against our vesicle preparations being artifacts of the
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propensity of lipids for self-assembly in solution.
Accepted Article
307
313
Vesicles were isolated from both planktonic cultures and bacterial biofilms. Biofilm is
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considered the primary state in which bacteria exist in the environment therefore it is perhaps not
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surprising that we identified extracellular vesicles within the matrix of B. subtilis biofilm.
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Vesicles isolated from all biofilm preparations were similar in size to planktonic vesicles,
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implying natural vesicle production by B. subtilis despite different growth conditions. Notably,
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vesicles isolated from strain 168 biofilm were free floating and non-adherent to other material
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whereas strain 3610 biofilm vesicles were often associated to flagella and other debris. This may
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reflect reduced production of exopolysaccharide by strain 168 due to a mutation in the epsC gene
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(McLoon et al., 2011).
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Previously, Tashiro et al showed that OMVs with stronger ζ potentials were more likely
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to associate with cells in P. aeruginosa (Tashiro et al., 2010). B. subtilis cells have a strongly
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negative ζ potential with a magnitude similar to the vesicles recovered from the supernatant.
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This finding is consistent with notion that B. subtilis extracellular vesicles originate from the cell
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membrane and have the potential to interact with bacterial cells (Ahimou et al., 2001).
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When isolating vesicles from the B. subtilis 168 lab and 3610 environmental strains, we
328
noted that vesicles isolated from each strain under the same culture conditions resulted in
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different DLS profiles. Strain 168 produced two populations similar in size to vesicles from
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other organisms whereas strain 3610 showed signals not typically associated with vesicles and
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more consistent with the presence of contaminating structures. The apparent paucity of vesicle
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signatures in the DLS profile of vesicle preparation from strain 3610 most likely reflects flagella
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contamination, which could skew the DLS data. For strain 168, vesicles were so abundant
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relative to any contaminating flagella that they exhibited a profile like those reported for other
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vesicle preparations (Rivera et al., 2010). DLS results of sfp+ strains, SL3610 and RL3090, also
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demonstrated similar contaminant distributions to strain 3610 whereas strain AD3610 (sfp-)
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displayed two clean peaks like strain 168. The dimensions obtained from DLS data were
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consistent with measurements from TEM images, which also revealed that more vesicles could
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be recovered from strain 168 compared to strain 3610. Strain 168 vesicle preparations contained
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so many vesicles that they were identified in the entire EM field of view whereas in strain 3610
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vesicle preparations, only an occasional vesicle was present in microscopic fields. It is notable
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that cells from all strains exhibited putative vesicular blebs on the cell membrane in SEM images
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even though we recovered various amounts of vesicles from the strains (data not shown).
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Although such blebs cannot be conclusively identified as vesicles based on sharing similar
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morphology and dimensions, their presence in the surface of wild type strains, and their
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relatively scarcity in the culture supernatants of those strains is consistent with, and supportive of
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the notion that they are made by wild type strains but disrupted by surfactin after release from
348
cells.
Accepted Article
329
349
The ability of strain 168 to be easily genetically manipulated as well as to produce large
350
quantities of vesicles makes it an ideal model strain to study the genetics and mechanisms of
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vesicle production in Gram-positive bacteria. Further, mutations in sfp, epsC, swr,and PdegQ, as
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well as the lack of the plasmid-borne gene rapP in strain 168 were identified as genetic
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differences between strain 168 and strain 3610 (McLoon et al., 2011). We further investigated
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the effect of the sfp gene on vesiculogenesis. Strain AM373 (168 harboring wild type copies of
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aforementioned genes) showed decreased amounts of recoverable vesicles compared to the
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background 168 strain by TEM and metabolic labeling (data not shown). This was evidence that
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the lack of a functional copy of one or more of these genes could be responsible for high vesicle
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recovery in the 168 strain. We investigated the candidate gene, sfp, further for its effects on
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vesicle production in B. subtilis. When a functional copy of sfp was restored into strain 168, the
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quantity of recoverable vesicles was greatly reduced (RL3090). Furthermore, when sfp was
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disrupted in strain 3610 (AD3610), vesicle recovery levels increased similar to strain 168. Strain
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SL3610, a sfp complement strain of AD3610, again showed abrogated vesicle recovery. These
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results suggest that sfp is an important factor in the amount of vesicles recovered between wild
364
strains and lab strains of B. subtilis and further reinforces the authenticity of vesiculogenesis by
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the Gram-positive bacterium.
366
recently been reported in mycobacteria (Rath et al., 2013).
Accepted Article
352
We note that genetic regulation of vesicular production has
367
Sfp protein is required for surfactin biosynthesis; therefore strains lacking sfp cannot
368
produce surfactin. Surfactin has been shown to disrupt lipid membranes and so we investigated
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surfactin as the mechanism behind recovering variable amounts of vesicles between wild and
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domestic strains of B. subtilis. We were able to recover similar amounts of vesicles from strain
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RL2663 as from strain 168. RL2663 has a functional sfp gene but a mutation in srfAA, which is,
372
also required for surfactin synthesis. This data led us to believe that surfactin production affected
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the amount of recoverable vesicles. We confirmed that vesicle incubation in pure surfactin and
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cell-free supernatant from surfactin-producing strain 3610 leads to less recoverable radioactivity
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in the vesicle pellet compared to incubation in surfactin-nonproducing strain 168 supernatant,
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PBS, BHI broth, and 10% EtOH indicative of vesicle disruption. We also determined that strain
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3610 supernatant and pure surfactin disrupted vesicles from B. anthracis, suggesting that wild B.
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subtilis can utilize surfactin to disrupt extracellular vesicles produced by other species. The high
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stability of B. anthracis vesicles in strain 168 supernatant, which corresponds to depleted BHI
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media, may be due to the cells utilizing one or more destabilizing molecules in the media but this
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difference was not pursued further.
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functions as iron transport, it is conceivable that surfactin-mediated disruption of vesicles from
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other species provides a competitive advantage to B. subtilis (Rath et al., 2013). In summary,
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our data implies that environmental strains produce vesicles that are disrupted by surfactin and
385
provides a new function for this lipopeptide in promoting release of vesicular cargo into the
386
extracellular space.
Accepted Article
375
Since vesicles have been implicated in such critical
387
In conclusion, we have characterized extracellular vesicles produced by Bacillus subtilis.
388
These vesicles are not only similar in size and morphology to other bacterial vesicles but we
389
were also able to describe vesiculogenesis in various strains. In addition, the ability to recover a
390
large amount of vesicles from strain 168 makes it an efficient model for the study of
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vesiculogenesis in Gram-positive bacteria. We have identified a gene, sfp, which is
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nonfunctional in most laboratory strains, that indirectly affects the quantity of vesicles
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recoverable from cell cultures. The lipopeptide surfactin, which requires functional sfp, disrupts
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vesicles in the extracellular milieu to release vesicle cargo. To our knowledge, this is the first
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time a mechanism for self-and foreign-vesicle cargo release has been reported in Gram-positive
396
bacteria. The occurrence of vesicular production in B. subtilis combined with the ability to
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induce release of vesicular content across species boundaries provides a potentially outstanding
398
platform for the dissection of this enigmatic process in a model organism.
399
Experimental Procedures
400
Bacterial Strains
Accepted Article
397
401
Bacillus subtilis 168 strain was obtained from ATCC #23857, and B. subtilis strains
402
NCIB# 3610, AMA373, RL3090, RL2663, and AD3610 were gifts from Richard Losick
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(Boston, MA) (Table 1). Strain AD3610 was constructed using techniques previously described
404
(Yasbin; Young, 1974; Gryczan et al., 1978; Wach, 1996). Briefly, PCR was utilized to disrupt
405
the sfp gene with a Kan cassette (Table S1). The construct was transformed into strain PY79 and
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moved into strain 3610 by phage transduction. Strain SL3610 was constructed by ectopic
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integration of sfp-spec into the AmyE locus of AD3610 by using pDG1730 and In-Fusion HD
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Cloning Plus kit (Clontech) (Table S1). Transformation was performed as stated above. The lab
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strain of B. anthracis Sterne 34F2 (pXO1+, pXO2−) used in stability assays was obtained from
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Alex Hoffmaster at the Center for Disease Control (Atlanta, GA). All strains were grown and
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maintained from 25% glycerol -80oC frozen stock on brain heart infusion (Difco) agar or broth at
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37oC. Antibiotics used include 0.5 μg ml-1 erythromycin and 2.5 μg ml-1 lincomycin (mls), 100
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μg ml-1 spectinomycin, and 10 μg ml-1 kanamycin.
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Vesicle Isolation
415
Vesicles were obtained with minor modifications of the method used previously for the
416
isolation of B. anthracis vesicles (Rivera et al., 2010). Briefly, bacterial cultures were spun for
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20 min at 4oC at 15,000 x g to remove cells. The resulting supernatant was then filtered through
418
a 0.22 um Milipore Express Plus Membrane and the filtrate was concentrated using Amicon
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ultrafiltration system (Millipore) with 100 kDa filter. The concentrate was then further filtered
420
through a 0.8 um syringe filter (Corning) to remove larger cellular debris or aggregated material.
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The filtered supernatant was then centrifuged at 195,000 x g for 1 h at 4oC and washed with
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phosphate buffered saline (PBS). The vesicle pellet was resuspended in PBS or water. Heat-
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killed culture vesicles were prepared by autoclaving an overnight culture. The cells were then
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spun for 20 min at 4oC at 15,000 x g and washed with PBS. The dead cells were then suspended
425
in fresh media and incubated overnight with shaking. Vesicles were then isolated in parallel with
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live cultures.
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Vesicle Size by Dynamic Light Scattering (DLS)
Accepted Article
419
428
The size distribution and diameter of vesicles resuspended in PBS or water was measured
429
using 90Plus/BI-MAS Multi Angle Particle Sizing analyzer (Brookhaven Instruments Corp.), as
430
previously described (Eisenman et al., 2009).
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Transmission Electron Microscopy (TEM)
432
Vesicles were visualized by TEM using negative and positive staining techniques. For
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negative staining, vesicles were adhered to 300-mesh carbon and Formvar coated Copper grids
434
for 2 min, stained with 1% phosphotungstic acid and viewed on a JEOL 100CXII or 1200EX at
435
80kV.
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paraformaldehyde in 0.1 M sodium cacodylate buffer. Samples were then postfixed with 1%
437
osmium tetroxide followed by 2% uranyl acetate en-bloc staining. The samples were dehydrated
438
through a graded series of ethanol and embedded in LX112 resin (LADD Research Industries,
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Burlington VT). Pellicle biofilms were fixed for 24 h with 2.5% gluteraldehyde, 0.2 M sodium
440
cacodylate, 0.4 M sucrose, and 10mM MgCl2 (pH 7.4) in a 1:1 ratio with the MSgg culture media.
For embedded TEM, vesicles were fixed with 2.5% glutaraldehyde and 2%
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Pieces of each fixed biofilm were processed in suspension on a rotator. The biofilm samples were
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postfixed with 1% osmium tetroxide, 0.7% potassium ferrocyanide for 1 h followed by 1% uranyl
443
acetate. The biofilm samples were dehydrated through a graded series of ethanol and then
444
embedded in LX112 resin (LADD Research Industries, Burlington VT). Ultrathin (80 nm) sections
445
of all samples were cut on a Reichert Ultracut UCT, stained with uranyl acetate followed by lead
446
citrate.
447
Scanning Electron Microscopy (SEM)
Accepted Article
441
448
Cells were fixed in 5% glutaraldehyde, 0.2 M sodium Cacodylate, 0.4 M Sucrose, 10mM
449
MgCl2 pH 7.4 and mixed 1:1 with suspension. The biofilm samples were postfixed for 30 min with
450
1% osmium tetroxide, 0.7% potassium ferrocyanide, 0.1 M sodium cacodylate, 0.2 M sucrose, and
451
5 mM MgCl2 (pH 7.4). Samples were dehydrated through a graded series of ethanol and critical
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point dried using liquid carbon dioxide in a Tousimis Samdri 795 Critical Point Drier (Rockville,
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MD). The samples were sputter coated with gold-palladium in a Denton Vacuum Desk-2 Sputter
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Coater (Cherry Hill, NJ). Prepared cells were then examined in a Zeiss Supra Field Emission
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Scanning Electron Microscope (Carl Zeiss Microscopy, LLC North America), using an accelerating
456
voltage of 5 KV.
457
Glucuronoxylomannan Immunogold TEM
458
The polysaccharide, glucuronoxylomannan (GXM), was isolated from Cryptococcus
459
neoformans and fractionated to less than 10 kDa in size by filtration through Amicon Ultra
460
Centrifugal Filters (Millipore). GXM was added to a culture of strain 168 at inoculation and
461
incubated at 37oC for 18 h. Vesicles were then isolated and either sonicated (20 sec sonication
462
and 30 sec incubation on ice, repeated 4 times) or not (Sonic Dismembrator Model 100; Fisher
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Scientific). Vesicles were fixed with 2% paraformaldehyde, 0.1 % glutaraldehyde in 0.1 M
464
Phosphate buffer, followed by dehydration with a graded series of ethanol, and a progressive
465
lowering of the temperature to -20° in a RMC FS 7500. Samples were then embedded in
466
Lowicryl HM-20 monostep resin (Electron Microscopy Sciences) and polymerized using UV
467
light. Ultrathin sections were cut on a Reichert Ultracut UCT.
Accepted Article
463
468
Immunogold labeling was performed by fixing the samples in 4% paraformaldehyde
469
(Electron Microscopy Sciences), 0.1% glutaraldehyde (Polysciences), and buffered in 0.1 M sodium
470
cacodylate (pH 7.2). Samples were incubated with the primary monoclonal antibody (mAb) 18B7
471
against GXM diluted in incubation buffer overnight at 4oC. The sample was washed with BSA-C
472
(Electron Microscopy Sciences) and incubated with 10 nm immunogold-conjugated secondary
473
antibody (donkey anti-mouse) (Electron Microscopy Sciences) for 2 h at room temperature.
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Samples were washed again and postfixed with 2% gluteraldehyde/PBS. Finally, samples were
475
stained with 4% uranyl acetate and imaged using JEOL 100CXII transmission electron microscope
476
at 80kv. Controls included preparations with no primary antibody as well as preparations with an
477
irrelevant IgG antibody.
478
Metabolic labeling with C-14 Palmitic Acid
479
B. subtilis and B. anthracis cells were labeled with radioactivity as previously described
480
for C. neoformans with minor modifications (Wolf et al., 2012). Briefly, an overnight culture of
481
bacteria was diluted to a suspension with an optical density at 600 nm (OD600) of ~0.1. 1-2 μCi
482
of 14C PA was added to aliquots of 15-30 ml. The samples were incubated at 37oC until OD600
483
doubled to ~0.2 and the cells were then pelleted, washed, and resuspended in fresh BHI.
484
Aliquots of 1 ml were added to 14 ml BHI in PETG flasks (Thermo Scientific) and grown
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overnight at 37oC. Overnight cultures were normalized by OD600 and vesicles were harvested by
486
centrifuging cells for 20 min at 15,000 x g at 4 oC and filtering the supernatant with a 0.8 μm
487
syringe filter (Corning) and the resultant supernatant was concentrated with Amicon Ultra
488
Centrifugal Filters 100 kDa (Millipore). The concentrate was spun at 195,000 x g for 1 h at 4 oC,
489
and washed with PBS. The vesicle pellet was then resuspended in PBS for dynamic light
490
scattering and scintillation counting (CPM). Controls included B. subtilis incubated without 14C
491
PA, cells incubated with 150 mM NaN3, and cultures without B. subtilis cells.
492
production over time was studied by counting radioactivity of vesicles isolated at 0, 4, 12, and 18
493
h. Samples measured include cell pellets, supernatant, concentrate FT, concentrate supernatant,
494
and vesicle pellet and each fraction represented as a ratio of the total sample. Scintillation
495
counting was done in a Packard Bioscience Tricarb A2900 liquid scintillation counter. The
496
position of vesicle fractions was determined by resuspending labeled vesicles in 30% Optiprep
497
Density Gradient Medium (Sigma) and layering 2 ml of 25, 20, 15, and 10% Optiprep above the
498
vesicle sample. The gradient was spun for 16 h at 140,000 x g at 4oC. 0.5 ml fractions were
499
taken from the top of the gradient and counted.
500
Vesicle Stability
Accepted Article
485
501
Vesicle stability over time, incubated in PBS, was measured by incubating
Vesicle
14
C PA-
502
labeled vesicle pellets at room temperature for 0, 24, 48 and 72 h in PBS followed by
503
ultracentrifugation and counting of supernatants and pellets. Vesicle stability was measured in
504
10 μg ml-1 and 100 μg ml-1 surfactin (resuspended in EtOH at 10 mg ml-1) (Sigma), BHI media,
505
PBS, 10% EtOH, and cell-free supernatants from B. subtilis strains 168 and 3610 week old
506
cultures. B. anthracis stability was measured similarly to B. subtilis but with the added condition
507
of cell-free supernatant from B. anthracis strain Sterne 34F2. Vesicle stability was measured by
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14
C PA-labeled vesicle pellets resuspended in the various conditions at 37 oC on a
incubating
509
nutator and after 24 h, samples were ultracentrifuged and radioactivity of supernatants and
510
pellets were counted.
511
ζ Potential Measurements
Accepted Article
508
512
The ζ potential of vesicles in water with 10mM dextrose and 1mM KCl was calculated
513
using a ζ potential analyzer (ZetaPlus; Brookhaven Instruments Corp.).
514
Protein Characterization
515
Vesicles were isolated and purified by density gradient centrifugation using OptiPrep and
516
vesicle containing fractions were dialyzed overnight against PBS at 4oC with one buffer change.
517
Cells were grown for 18 h at 37o, spun down and washed with PBS. Cell membrane fractions
518
were isolated as previously described (Petrackova et al., 2010).
519
lyophilized and analyzed.
All samples were then
520
Proteins were precipitated from vesicle suspensions, membrane fractions, and bacterial
521
lysates by adding six volumes of ice-cold acetone. After incubation overnight, the acetone was
522
removed and the pellets dissolved in 25 mM ammonium bicarbonate. Proteins were reduced with
523
10 mM DTT at room temperature for 1 h and the cysteine residues were subsequently alkylated
524
with 10 mM iodoacetamide at room temperature for 1h in the dark. Protein digestion was carried
525
out with trypsin (Promega, 1:20, w/w) at 37ºC overnight. The peptide mixtures from in-solution
526
tryptic digestions were analyzed using nanoflow LC-MS/MS. The peptides were loaded onto a
527
0.3 x 5 mm C18 precolumn (New Objective) and then eluted with a linear gradient of 2-90%
528
acetonitrile in 0.1% aqueous solution of formic acid. The gradient was delivered over 120 min by
529
a nanoLC-1Dplus (Eksigent) at a flow-rate of 200 nl/min, through a 75-µm x 15 cm fused silica
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capillary C18 HPLC PepMap column (LC Packings) to a stainless steel nano-bore emitter
531
(Proxeon). The peptides were scanned and fragmented with an LTQ XL linear ion trap mass
532
spectrometer (ThermoFinnigan) operated in data-dependent and MS/MS switching mode using
533
the three most intense precursors detected in a survey scan from 400 to 1600 amu (three µscans).
534
Normalized collision energy was set to 35%, and dynamic exclusion was applied during 3 min
535
periods to avoid repetitive fragmentation of ions. Generated .raw files were converted to .dta
536
files for Mascot database. A database containing the NCBI B. subtilis (4176 sequences, as of
537
January 20, 2012) was searched using Mascot Software (version 2.3, Matrix Science) for protein
538
identification. Search criteria included trypsin specificity with one missed cleavage allowed, and
539
methionine oxidation as variable modification. A minimum precursor and fragment-ion mass
540
accuracy of 1.2 and 0.3 Da, respectively, and a requirement of at least one bold red peptide (i.e.
541
highest scoring peptide matching to protein with highest total score). Cut-off values for Mascot
542
scores of peptides and proteins were set to 45 (p < 0.05) and 51 (p < 0.01), respectively, for
543
considering them as being accurate identifications. Proteins identified with only one peptide
544
were inspected manually. The false positive rate (1,23%) was calculated searching the same
545
spectra against the NCBI B. subtilis decoy database, using the version updated January 20, 2012.
546
A combined list of proteins identified in all replicates was condensed in order to remove
547
redundant IDs such as orthologous sequences, redundant database entries, and indistinguishable
548
isoforms based on observed peptide coverage.
Accepted Article
530
549
Vesicle preparation protein content from strains 168, 3610, and RL2663 was normalized
550
with Pierce BCA Protein Assay Kit (Thermo Scientific) and 7.5 ug of protein was run on 4–20%
551
Mini-PROTEAN® TGX Stain-Free Gel (Bio-Rad) and silver stained using Pierce Silver Stain
552
Kit (Thermo Scientific).
This article is protected by copyright. All rights reserved.
Biofilm Assay and Vesicle Isolation
Accepted Article
553 554
Biofilms were grown as described in McLoon et al. with modifications (McLoon et al.,
555
2011). Bacteria were grown from frozen stock on BHI agar overnight at 37oC. A single colony
556
was then inoculated into BHI broth for 4 h at 37oC with shaking. The culture was inoculated
557
(1:1000) into MSgg media (100 mM MOPS, pH 7, 5 mM potassium phosphate, pH 7, 700 μM
558
CaCl2, 2 mM MgCl2, 50 μM FeCl3, 50 μM MnCl2, 1 μM ZnCl2, 2 μM thiamine, 50 μg ml-1
559
tryptophan, 50 μg ml-1 phenylalanine, and 50 μg ml-1 threonine, 0.5% glycerol and 0.5%
560
glutamate). Biofilms were grown at 30oC for either 4 or 8 d without shaking. After the desired
561
time, the biofilm culture was disrupted and centrifuged at 15,000 x g for 15 min to remove
562
cellular debris. The resulting supernatant was filtered through a 0.22 um Milipore Express Plus
563
Membrane and then concentrated using Amicon Ultra Centrifugal Filters 100 kDa (Millipore).
564
The resultant concentrate was centrifuged at 195,000 x g at 4 oC and washed with PBS. The
565
vesicle pellet was resuspended in PBS or water. Whole biofilm pellicles were submitted to TEM
566
and SEM preparations.
567
Statistics
568
Statistics were performed in Prism (GraphPad Software). Error bars indicate mean and
569
standard deviation. Comparisons between ratio of natural and sonicated vesicles containing gold
570
particles was done using Chi2 test.
571
compared using an unpaired t-test. One-way ANOVA was performed comparing vesicle density
572
and diameter. Vesicle pellet radioactivity between strains 168 and 3610 was analyzed with an
573
unpaired t-test. A one-way ANOVA was utilized when comparing vesicle pellet radioactivity
574
among strains. One-way ANOVA was also used to compare stability of pellet radioactivity after
The diameters of natural and sonicated vesicles were
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incubation in various conditions. Significance was indicated as follows: p
