Bacteria are known to communicate primarily via secreted extracellular factors. Here we identify a previously uncharacterized type of bacterial communication mediated by nanotubes that bridge neighboring cells. Using Bacillus subtilis as a model organism, we visualized transfer of cytoplasmic fluorescent molecules between adjacent cells. Additionally, by coculturing strains harboring different antibiotic resistance genes, we demonstrated that molecular exchange enables cells to transiently acquire nonhereditary resistance. Furthermore, nonconjugative plasmids could be transferred from one cell to another, thereby conferring hereditary features to recipient cells. Electron microscopy revealed the existence of variously sized tubular extensions bridging neighboring cells, serving as a route for exchange of intracellular molecules. These nanotubes also formed in an interspecies manner, between B. subtilis and Staphylococcus aureus, and even between B. subtilis and the evolutionary distant bacterium Escherichia coli. We propose that nanotubes represent a major form of bacterial communication in nature, providing a network for exchange of cellular molecules within and between species.

Tubular conduits between cells that allow exchange of cellular content are typical of multicellular organisms. In plants, neighboring cells are connected by cytoplasmic tubes called plasmodesmata, which provide multiple routes for intercellular transfer of nutrients, signals, proteins and transcripts (). In mammalian cells, intercellular communication is mediated locally through gap junctions and synapses; however, recent reports demonstrate the existence of a network of intercellular membrane nanotubes enabling long-distance communication. These tunneling nanotubes have been shown to facilitate intercellular transfer of cytoplasmic molecules and even organelles and viruses (). Here we report the identification of analogous nanotubular channels formed among bacterial cells grown on solid surface. We demonstrate that nanotubes connect bacteria of the same and different species, thereby providing an effective conduit for exchange of intracellular content.

An additional type of molecular exchange that involves physical interactions between neighboring bacterial cells is conjugation (). During this process DNA is transferred from a donor to a recipient through a pilus, a tube-like structure that physically connects the participating cells (). Notably, conjugation represents a key mechanism of horizontal gene transfer in nature (), whereby hereditary genetic information, rather than nonhereditary molecular signal, is delivered.

Secretion and detection of small extracellular molecules to the surrounding environment is not the only form of molecular exchange between bacteria. Many Gram-negative bacteria trade information by packaging molecules into extracellular membrane vesicles (MVs). These MVs can travel and fuse with distal cells, thus providing a secure mode for delivering various cellular moieties, including QS molecules, antimicrobial factors, toxins, and DNA (). Furthermore, in some cases neighboring daughter cells have been found to exchange molecular information by establishing intimate cytoplasmic connections. In cyanobacteria, for example, the movement of small molecules (e.g; sugars and amino acids) within a filament was shown to be mediated by intercellular channels. This cytoplasmic sharing enables vital cooperative behavior between nitrogen fixing heterocysts and photosynthetic nurturing cells ().

Multicellular activity is achieved by the ability of group members to exchange information in order to synchronize their behavior. Importantly, bacteria are not limited to communicate within their own species but are also capable of sending and receiving messages in an interspecies manner. In both Gram-positive and -negative bacteria, cell-to-cell exchange of information is mediated primarily by signaling molecules belonging to the general classes of low molecular weight autoinducers and signaling oligopeptides (). In a process known as quorum sensing (QS), the production and detection of these signaling molecules is employed by bacteria to monitor population density and modulate gene expression accordingly ().

Bacteria in nature display complex multicellular behaviors that enable them to execute sophisticated tasks such as antibiotic production, secretion of virulence factors, bioluminescence, sporulation, and competence for DNA uptake (). Such social activities ultimately benefit the population and are unproductive if performed by a single bacterium. Furthermore, nearly all bacteria are capable of forming a resilient multicellular structure, termed biofilm, comprising cells with different functionalities. Natural biofilms are typically composed of several bacterial species and therefore demand a coordinated gene expression of the various inhabitants ().

Finally, we addressed whether cytoplasmic molecules can be traded between evolutionary distant Gram-positive and -negative bacteria. We therefore tested the capability of B. subtilis cells to transfer cytoplasmic molecules to the Gram-negative bacterium Escherichia coli. When B. subtilis (gfp+) cells were cultured alongside E. coli (gfp−) cells, a pronounced fluorescence gradient developed in a temporal manner ( Figure 7 A and Figure S3 Bc). Accordingly, interspecies nanotubes directly bridging neighboring B. subtilis and E. coli cells were evident by HR-SEM ( Figures 7 B and 7C). In general, the nanotubes formed by E. coli cells appeared significantly thinner than those formed by B. subtilis or S. aureus, suggesting that Gram-positive and -negative bacteria form somewhat different types of nanotubes. HR-SEM also clearly revealed the presence of nanotubes formed between S. aureus and E. coli ( Figure 7 D) demonstrating the ubiquitous nature of this phenomenon.

(D) S. aureus (MRSA) and E. coli (MG1655) cells were grown to midexponential phase. Grown cells were mixed (1:1 ratio), plated on LB agar, incubated for 6 hr at 37°C, and visualized by HR-SEM (×75,000). An arrow indicates interspecies tubes connecting two neighboring cells. The scale bar represents 250 nm.

(B and C) B. subtilis (PY79) and E. coli (MG1655) cells were grown to midexponential phase. Grown cells were mixed (1:1 ratio), plated on LB agar, incubated for 6 hr at 37°C, and visualized by HR-SEM (see Experimental Procedures ). (B) A typical field of the mixed population (×100,000) is shown. The red circle highlights nanotubes between neighboring B. subtilis and E. coli cells. (C) A higher-magnification image (×400,000) of the circled region in (B). Based on texture similarity, the green arrow denotes a thick nanotube emanating from the B. subtilis cell and the blue arrow indicates a thinner nanotube emanating from the E. coli cell. The scale bars represent 500 nm (B) and 200 nm (C).

(A) Exponentially growing cells of B. subtilis SB444 (gfp+) and E. coli (MG1655) (gfp−) strains were mixed (1:1 ratio), plated on an LB agarose pad, and incubated in a temperature controlled chamber at 37°C. Cells were visualized by time-lapse fluorescence microscopy, and images of phase contrast (blue) and fluorescence (green) were collected at 10 min intervals. Select overlay (a–c) and GFP (a′–c′) images are shown from the following time points: (a and a′) t0 min (b and b′), t30 min, and (c and c′) t50 min. The scale bar represents 1 μm.

Examining a field of cocultured cells by HR-SEM revealed visible intercellular bridges among the cells of each species, but, more importantly, clear protrusions were formed between species ( Figure 6 B). Visualizing the intercellular nanotubes formed between S. aureus cells in high resolution revealed morphology and dimensions similar to the large tubes formed by B. subtilis cells ( Figure 6 C). The interspecies B. subtilis-S. aureus connections also displayed morphology resembling that of B. subtilis nanotubes ( Figure 6 D and Figure S4 H). We infer that Gram-positive bacteria can trade cellular information within and between species by a path comprising of the intercellular connections.

To broaden our investigation, we examined whether the exchange of cytoplasmic molecules and the formation of intercellular nanotubes occur between species. First, we investigated the ability of B. subtilis cells to transfer cytoplasmic molecules to the Gram-positive coccus, Staphylococcus aureus. B. subtilis (gfp+) and S. aureus (gfp−) cells were cocultured and followed by time-lapse fluorescence microscopy. Remarkably, at t30 min, a significant fluorescence signal was acquired by the S. aureus cells in a manner proportional to their distance from the gfp+ bacilli cells ( Figure 6 A ). This phenomenon was reinforced after 50 min of coincubation ( Figure 6 A and Figure S3 Bb). We surmise that molecular transfer can take place between two distinct species of Gram-positive bacteria that reside in proximity.

(B–D) B. subtilis (PY79) and S. aureus (MRSA) cells were grown to midexponential phase. Grown cells were mixed (1:1 ratio), plated on LB agar, incubated for 6 hr at 37°C, and visualized by HR-SEM (see Experimental Procedures ). (B) A typical field of the mixed population (×12,000). Blue arrows indicate visible intercellular nanotubes between S. aureus cells, whereas green arrows point to interspecies B. subtilis and S. aureus connecting tubes. (C) A high-magnification image (×50,000) of a nanotube connecting two S. aureus cells. (D) A high-magnification image (×50,000) of an interspecies nanotube connecting S. aureus and B. subtilis cells. The scale bars represent 5 μm (B) and 0.25 μm (C and D).

(A) Exponentially growing cells of B. subtilis SB444 (gfp+) and S. aureus (MRSA) (gfp−) strains were mixed (1:1 ratio), plated on an LB agarose pad, and incubated in a temperature controlled chamber at 37°C. Cells were visualized by time-lapse fluorescence microscopy, and images of phase contrast (blue) and fluorescence (green) were collected at 10 min intervals. Select overlay (a–c) and GFP (a′–c′) images are shown from the following time points: (a and a′) t0 min (b and b′) t30 min and (c and c′) t50 min. The scale bar represents 1 μm.

We exploited the antibiotic resistance assay to screen for bacterial genes that influence nanotube formation. Specifically, we analyzed an array of mutants in cell division, cell shape and membrane metabolism for their capacity to exchange antibiotic resistance ( Table S1 ). However, none of the tested mutants significantly reduced the antibiotic resistance of the mixed population. It is possible that nanotube production is induced by several overlapping mechanisms involving different gene families in a cooperative manner.

To examine the SDS sensitivity of nanotubes, cells grown in the presence of SDS were visualized with HR-SEM. Indeed, when we observed cells growing at 0.007% SDS, a concentration in which acquiring antibiotic resistance was clearly decreased ( Figure S7 A), only few intercellular nanotubes could be discerned ( Figure S7 C). However, prominent nanotubes were evident in untreated cells ( Figure S7 D). These SDS experiments indicate that nanotubes are composed of membrane components and further correlate the integrity of nanotubes with the exchange of antibiotic resistance among neighbors.

Next, we asked if, similar to their eukaryotic counterparts and as indicated by our TEM analysis, bacterial nanotubes are indeed composed of membrane constituents ( Figure S5 ). To explore this possibility, we examined nanotubes sensitivity to the membrane detergent sodium dodecyl sulfate (SDS). Initially, we tested whether SDS influences the phenomenon of acquiring nonhereditary antibiotic resistance from neighboring cells. Hence, cocultured P1 (Cm) and P2 (Kan) cells were spotted onto LB plates containing different concentrations of SDS, and their ability to grow on Cm+Kan plates was assayed by replica plating. An inverse correlation was observed between growth of the mixed cells on the Cm+Kan plate and SDS concentration ( Figure S7 A ). Importantly, at a concentration of 0.009% SDS, the ability of the cells to grow on the Cm+Kan selective plate was abolished, yet their viability was not significantly affected ( Figure S7 B). These results show that SDS prevents acquisition of antibiotic resistance from nearby cells, suggesting bacterial nanotubes are SDS sensitive.

(D) PY79 cells were grown to midexponential phase as in (C) in the absence of SDS and visualized by HR-SEM. Shown is a typical field of cells whereby a network of intercellular connecting nanotubes is evident (×10,000). The scale bar represents 5 μm.

(C) Wild-type PY79 cells were grown to midexponential phase, plated on LB agar containing 0.007% SDS, incubated for 6 hr at 37°C and visualized by HR-SEM (see Experimental Procedures). Shown is a typical field of cells evidently lacking visible nanotubes (×10,000). The scale bar represents 2 μm. In general, cells grown in the presence of SDS appeared in long chains and frequently failed to stick to the EM grids properly. This made it difficult to visualize cells growing at higher SDS concentrations.

(B) P1 and P2 strains were grown to midexponential phase and spotted in serial dilutions (3 −6 -3 −10 ) on LB agar containing SDS at the indicated concentrations. To rule out the possibility that SDS increases antibiotic sensitivity, plates were supplemented with Cm for P1 and Kan for P2. Cells were photographed after O/N. In general, cells spotted on SDS containing plates spread more in diameter than cells spotted on plates without SDS. On high SDS concentrations cell growth was delayed; however, as evidence here, the viability was not significantly reduced.

(A) An antibiotic assay examining the transfer of Cat protein under different SDS concentrations. Left: equal numbers of cells from PY79 (WT), SB463 (amyE::Phyper-spank-cat-spec) (P1: Cm R ) and SB513 (amyE::Phyper-spank-gfp-kan) (P2: Kan R ) strains were spotted separately on LB agar plates containing the indicated amounts of SDS. In parallel, equal numbers of mixed P1 and P2 cells (1:1) were spotted similarly. Spotted cells were grown for 4 hr at 37°C. Right: grown cells were replica plated onto the indicated plates supplemented with SDS. Plates were incubated O/N at 37°C.

We conclude that when grown on solid surface, B. subtilis cells are able to exchange nonconjugative plasmids. Unlike conjugation, which is induced by genes carried on conjugative plasmids, intercellular nanotubes may provide a constitutive path for reciprocal genetic exchange in nature without the need for a donor or a recipient strain.

To substantiate that the plasmid was not delivered to recipient cells by transformation, P2 cells were incubated with an excess amount of exogenous pHB201 DNA. Exogenous addition of plasmid DNA was unable to allow growth of P2 cells on the Cm+Kan plate, indicating that transformation is not the mechanism for plasmid exchange under our experimental conditions ( Figure 5 B). Furthermore, plasmid transfer was DNaseI resistant (see Extended Experimental Procedures and data not shown), implying that the transferred DNA is protected during passage from one cell to another by nanotubes that serve as a delivery vehicle. Similarly, DNaseI resistance was found to be a characteristic of conjugative plasmids passing through the protective conjugative tube (). Measuring the frequency of pHB201 transfer revealed a value of 10/colony forming unit (CFU) (see Extended Experimental Procedures ). In comparison, examining the transfer of a bona fide conjugative plasmid (pLS20) revealed a transfer frequency that was 1000 fold higher than pHB201 (10/CFU), similar to the frequencies reported previously ().

Given that nonhereditary features can be traded between nearby cells, we explored whether genetic information carried by an extrachromosomal plasmid can also be exchanged. To examine this possibility, we transformed B. subtilis strain P1 (SB463: amyE::Phyper-spank-cat-spec) (Cm, Spec) with a nonintegrative vector pHB201 (6.6 kb ∼4.35 MDa; cat, erm) (Cm, Mls) (). The resultant strain P1′ (GD110) was consequently Cm, Spec, and Mls(Mls is a mixture of Erm and Lin, both can be neutralized by erm). Next, P1 and P1′ were spotted separately or in a mixture with P2 strain (SB513: amyE::Phyper-spank-gfp-kan) (Kan, GFP+) on a nonselective plate, grown for 4 hr, and the antibiotic resistance of these populations was tested ( Figure 5 A ). Consistent with previous results, only cells within the mixed cultures (P1+P2 or P1′+P2) were able to grow on a plate containing both Cm and Kan ( Figure 5 A, first-replica plating). To distinguish exchange of nonhereditary molecules from plasmid delivery, we examined whether the observed dual resistance was heritable. Accordingly, cells growing on Cm+Kan were re-replica plated onto respective antibiotic plates to determine their genotype ( Figure 5 A, second-replica plating). In line with the data described above, cells from the P1+P2 mixture did not resume growth on the Cm+Kan plate but were able to grow on the Kan plate, implying that their dual resistance was a transient nonhereditary feature exhibited by recipient P2 cells. In contrast, a substantial fraction of the P1′+P2 population grew on the Cm+Kan plate and also on a plate containing Cm+Kan+Mls ( Figure 5 A, asterisk). The emergence of cells carrying Kan, Cm, and Mls resistances suggests that the plasmid pHB201 (Cm, Mls) was transferred from the donor P1′ (Kan) strain to the recipient P2 (Kan) strain. Indeed, these multiply resistant cells were all Specand GFP+, supporting the view that P2 (Spec, GFP+) rather than P1′ (Spec, GFP−) cells survived ( Figure 5 A, second-replica plating; data not shown). Finally, pHB201 could be extracted from the multiply resistant cells confirming that their phenotype was not a consequence of genetic mutations but derived from receipt of the extrachromosomal plasmid (data not shown).

(B) An antibiotic assay demonstrating that the plasmid is not transmitted by transformation. Left: (1) A mixture of GD110 (amyE::Phyper-spank-cat-spec, pHB201/cat, erm) (Cm R , Spec R , Erm R ) and SB513 (amyE::Phyper-spank-gfp-kan) (Kan R ) cells. (2–3) SB513 cells. Spotted cells were grown for 4 hr at 37°C in the presence or the absence of exogenous pHB201 DNA (100 ng of DNA/μl spotted cells) as indicated. Right: Grown cells were replica plated onto the indicated plates and incubated O/N at 37°C.

(A) An antibiotic assay examining the transfer of plasmids between B. subtilis cells. Left: Equal numbers of cells from P1 (SB463: amyE::Phyper-spank-cat-spec) (Cm R , Spec R ), P1′ (GD110: amyE::Phyper-spank-cat-spec, pHB201/cat, erm) (Cm R , Spec R , Mls R ), and P2 (SB513: amyE::Phyper-spank-gfp-kan) (Kan R ) strains were spotted separately on LB agar. In parallel, equal numbers of mixed P1+P2 (1:1) and mixed P1′+P2 (1:1) cells were spotted similarly. Cells were grown for 4 hr at 37°C. Right: Grown cells were replica plated onto the indicated plates (first-replica plating), and plates were incubated O/N at 37°C. Lower: To analyze the genotype of the cells growing on Cm+Kan antibiotic plate (highlighted with a green frame) cells were re-replica plated onto the indicated plates (second-replica plating). The plate labeled with an asterisk contains Cm+Kan+Mls. Plates were incubated O/N at 37°C.

Taken together, we conclude that the transient doubly resistant phenotype is a nonhereditary feature, and the resulting survivors are affected by the mechanism of antibiotic action.

Both Cm and Lin are bacteriostatic antibiotics that impede growth but do not instantly kill bacterial cells. On the other hand, bactericidal antibiotics kill bacteria rapidly, and thus necessitate constant protection by the resistance protein. Therefore, it remained possible that the dual resistance obtained by the mixed population would be affected if one of the participants harbors a gene imparting resistance to a bactericidal antibiotic. To examine this premise, we cocultured P1 strain (Cm) with P3 strain harboring chromosomally encoded resistance to the bactericidal antibiotic Kanamycin (Kan) and repeated the above assay ( Figure 4 C). In line with previous results, only cells in the mixed population were able to grow on the antibiotic plate containing both Cm and Kan ( Figure 4 C). Genotypic analysis of the surviving cells revealed that they were exclusively KanCm, implying that the P3 cells carrying the bactericidal antibiotic resistance gene survived ( Figure S6 B). Expanding this genotypic examination to thousands of colonies revealed that Cmcells (P1) survive rarely (∼1:700) under these conditions. This assay enables delineation between “donor” (Cm) and “recipient” (Kan) strains, providing an approach to follow the directionality of molecular exchange. To further confirm that the doubly resistant P3 cells indeed acquired Cat molecules from their neighbors, we carried out immunofluorescence microscopy with anti-Cat antibodies to detect the presence of Cat protein molecules within their cytoplasm (see Extended Experimental Procedures ). Consistent with our assumption, a clear fluorescent signal was detected from P3 cells grown in the mixture but was evidently absent from unmixed P3 cells ( Figure S6 C).

To test this prediction, we examined the exchange of chloramphenicol acetyltransferase (Cat) and erythromycin resistance methylase (Erm) between two different B. subtilis strains. The Cat protein confers resistance to chloramphenicol (Cm) and the Erm protein confers resistance to lincomycin (Lin). Strains harboring chromosomally encoded resistance to Cm (P1: Cm) or Lin (P2: Lin) were spotted separately or in a mixture onto LB agar plate and incubated for 4 hr in the absence of any antibiotic selection. Next, the ability of the strains to grow on selective plates containing Cm, Lin, or both was examined by replica plating ( Figure 4 B) in order to maintain the spatial arrangement of the cells. Strikingly, the mixed population of P1 and P2 cells was able to survive on the antibiotic plate containing both Cm and Lin ( Figure 4 B). To explore the genotype of the survivors, cells growing on the Cm+Lin plate were streaked onto a nonselective LB plate. Then individual colonies from the streak were picked, grown as stripes on LB plates, and their genotype was determined by replica plating onto Cm and Lin plates ( Figure S6 A ). Each tested colony exhibited either Cmor Lin, but not both, indicating that the surviving cells have not acquired a doubly resistant genotype. Summarily, we infer that nearby cells can exchange cytoplasmic molecules and gain transient nonhereditary phenotypes.

(C) Equal numbers of cells from P1 (SB463: amyE::Phyper-spank-cat-spec) (Cm) and P3 (SB513: amyE::Phyper-spank-gfp-kan) (Kan, GFP+) were spotted separately on LB agar. In parallel, equal numbers of mixed P1 and P3 cells were spotted similarly. Spotted cells were grown for 6 hr at 37°C. Grown cells were replica plated onto selective plates and finally onto LB as described in Figure 4 C. Replica plated P1 cells (grown on Cm plate), P3 cells (grown on Kan plate), and P1+P3 cells (grown on Cm+Kan plate), were scratched from the plates and subjected to immunofluorescence microscopy with anti-Cat antibodies (see Extended Experimental Procedures ). Left: Signal from anti-Cat antibodies (red), Middle: Signal from GFP (green), Right: Overlay of red and green signals with a phase contrast image (blue). Fluorescence images have been normalized to a similar intensity range. The scale bar represents 1 μm.

(B) An antibiotic assay examining the exchange of Cat and Kan resistance proteins (and possibly transcripts) between two different B. subtilis strains (see Figure 4 C). a: To analyze the genotype of the colonies growing on the Cm+Kan antibiotic plate shown in Figure 4 C, doubly resistant cells were streaked on LB plates and allowed to grow O/N. b: Single colonies were picked, streaked on LB plates and allowed to grow O/N. Parental strains (P1 and P3) were streaked as controls. c: The genotype of grown streaks was tested by replica plating onto Cm and Kan selective plates.

(A) An antibiotic assay examining the exchange of Cat and Erm proteins (and possibly transcripts) between two different B. subtilis strains (see Figure 4 B). a: To analyze the genotype of the colonies growing on the Cm+Lin antibiotic plate shown in Figure 4 B, doubly resistant cells were streaked on LB plates and allowed to grow O/N. b: Single colonies were picked, streaked on LB plates and allowed to grow O/N. Parental strains (P1 and P2) were streaked as controls. c: The genotype of the single colonies was tested by replica plating onto Cm and Lin selective plates.

Having established the existence of intercellular nanotube networks, we sought to explore their capability to generate new phenotypes. We anticipated that when two strains, each harboring a different antibiotic resistance gene, are grown together, the exchange of cytoplasmic molecules (proteins and possibly transcripts) through the tubes could yield a population of cells temporarily resistant to both antibiotics in a nonhereditary fashion ( Figure 4 A ).

(C) An antibiotic assay examining the exchange of Cat and Kan resistance proteins (and possibly transcripts) between two different B. subtilis strains. Left: Equal numbers of cells from PY79 (WT), SB463 (amyE::Phyper-spank-cat-spec) (P1: Cm R ), and SB513 (amyE::Phyper-spank-gfp-kan) (P3: Kan R ) strains were spotted separately on LB agar. In parallel, equal numbers of mixed P1 and P3 cells (1:1) were spotted similarly. Spotted cells were grown for 4 hr at 37°C. Right: Grown cells were replica plated onto the indicated selective plates and finally onto LB. Plates were incubated O/N at 37°C.

(B) An antibiotic assay examining the exchange of Cat and Erm proteins (and possibly transcripts) between two different B. subtilis strains. Left: Equal numbers of cells from PY79 (WT), SB463 (amyE::Phyper-spank-cat-spec) (P1: Cm R ), and GD57 (amyE::Phyper-spank-erm-spec) (P2: Lin R ) strains were spotted separately on LB agar. In parallel, equal numbers of mixed P1 and P2 cells (1:1) were spotted similarly. Cells were grown for 4 hr at 37°C. Right: Grown cells were replica plated onto the indicated selective plates and finally onto LB. Plates were incubated O/N at 37°C.

(A) A schematic model for the transient gain of nonhereditary phenotypes via intercellular nanotubes. Shown on the left are two B. subtilis cells, each harboring a different antibiotic resistance gene, providing Cm R or Lin R . Genes (colored stripes) are depicted on the chromosomes (olive lines) with colored circles and colored combs indicating their respective proteins and transcripts. Shown on the right is the gain of antibiotic resistance by proteins and transcripts passing through intercellular nanotubes in a mixed population. Molecular transfer through the connecting tubes yields a population of cells temporarily resistant to both antibiotics in a nonhereditary fashion.

To demonstrate that nanotubes indeed serve as a route for trading cytoplasmic molecules, we carried out immunoelectron microscopy (immuno-EM). gfp+ and gfp− cells were mixed and grown on solid medium. Next, cells were gently fixed, sectioned, incubated with anti-GFP antibodies and then immunostained with gold-conjugated secondary antibodies (see Extended Experimental Procedures ). Remarkably, the gold particles could be visualized within nanotubes connecting neighboring cells ( Figures 3 E–3G), corroborating that indeed intercellular nanotubes serve as a path for molecular exchange. In many images, a GFP gradient was observed whereby a GFP-producing cell containing multiple gold particles was connected to an adjacent cell containing few gold particles ( Figure 3 E), resembling the phenomenon observed by time-lapse microscopy ( Figure 1 C). Importantly, when only gfp− cells were similarly processed, no significant gold signal was detected (data not shown).

In an alternative approach, intercellular connections were visualized with transmission electron microscopy (TEM), where cells were imaged without employing any contrasting agent (see Extended Experimental Procedures ). Consistent with the HR-SEM images, a network of pronounced nanotubes tying one cell to another was readily visible ( Figure S5 A ). Interestingly, higher-magnification analysis of a typical tube appears to indicate a structure comprising outer and inner layers, hinting at a multilayered structure ( Figures S5 B–S5D). Moreover, thin section analysis suggests that the tubes contain cell wall material, membrane and cytoplasmic content ( Figures S5 E and S5F).

Tube dimension appears to vary with the distance between connected cells. Generally, tube length ranged up to 1 μm, whereas width ranged approximately from 30 to 130 nm (e.g., Figures S4 E and S4F). The relatively large size of the tubes concords with our assumption that they could easily accommodate the passage of proteins such as GFP (approximately 40 Å; []) and even larger cytoplasmic molecules. Closer investigation of the HR-SEM images revealed that beside the large nanotubes, an additional type of smaller nanotubes was visible, though more challenging to detect ( Figure 3 D). These smaller tubes tended to be clustered connecting nearby cells intimately, appearing to actually “stitch” one cell to another. We speculate that these smaller nanotubes are more ubiquitous than the larger ones and are capable of traversing small molecules.

The exchange of cytoplasmic molecules between adjacent cells raised the notion that intercellular connections, facilitating this process, exist. To examine this possibility, we grew B. subtilis cells (PY79) on solid Luria Bertani (LB) medium and visualized them with high-resolution scanning electron microscopy (HR-SEM; see Experimental Procedures ). Surprisingly, tubular protrusions (nanotubes) bridging neighboring cells were plainly visible ( Figure 3 A ). The nanotubes seem to project from the cell surface at different positions in a nonspecific manner. Higher-magnification micrographs clearly evidence a network of intercellular connecting nanotubes whereby cells frequently attach to more than one partner simultaneously ( Figures 3 B and 3C and Figure S4 A ). Occasionally, we observed the occurrence of branched nanotubes linking together several cells at once ( Figure S4 B). Notably, these tubes were structurally distinguishable from classical conjugative pili ( Figure S4 C). Examining cells of an undomesticated B. subtilis strain (3610) with the same procedure revealed a similar or an even enhanced ability to form nanotubes ( Figure S4 D). The existence of nanotubes was also detected when cells were incubated on minimal medium yet at a lower frequency ( Figure S4 G). However, nanotubes seem to be absent when cells were grown in liquid medium (data not shown), suggesting that growth on solid medium induces their formation.

(G) An additional example of an immuno-EM section, showing the localization of a GFP molecule within a tube, as indicated by an arrow. The scale bar represents 200 nm.

(E) An immuno-EM section of cocultured PY79 (gfp−) and SB444 (gfp+) cells, stained with anti-GFP and secondary gold-conjugated antibodies (see Extended Experimental Procedures ). Black dots indicate the expression and localization of GFP molecules. The scale bar represents 200 nm.

(A–D) PY79 cells were grown to midexponential phase, plated on LB agar, incubated for 6 hr at 37°C, and visualized by HR-SEM (see Experimental Procedures ). (A) A typical field of B. subtilis cells (×15,000). Green arrows indicate intercellular nanotubes connecting neighboring cells. The scale bar represents 5 μm. (B) A higher-magnification image (×40,000) of the boxed region in (A). Membrane bulging is indicated by an asterisk (). The scale bar represents 500 nm. (C) An additional field of cells demonstrating the occurrence of a network of intercellular nanotubes (×50,000). The scale bar represents 1 μm. (D) A field of cells where a cluster of smaller nanotubes (highlighted by a dashed circle) as well as a more pronounced larger tube (indicated by an arrow) are apparent (×100,000). The scale bar represents 500 nm.

Taken together, our results establish that adjacent B. subtilis cells are able to exchange cytoplasmic molecules in a spatially ordered manner. To the best of our knowledge, this is the first report of cytoplasmic sharing between neighboring B. subtilis cells.

In a complementary approach, cytoplasmic exchange was examined with calcein, a nongenetically encoded cytoplasmic fluorophore. Calcein is a small nonfluorescent acetoxymethylester (AM) derivative that is sufficiently hydrophobic to traverse cell membranes. After passage into the cytoplasm, hydrolysis of calcein by endogenous esterases gives rise to a fluorescent hydrophilic product (623 Da) unable to traverse membranes and thus caged within the cytoplasm (). When B. subtilis cells (PY79) were incubated with calcein-AM (see Extended Experimental Procedures ), they rapidly acquired a strong fluorescence signal indicating calcein hydrolysis. Next, labeled cells were washed, mixed with nonlabeled cells, and the mixture was placed on an agarose pad and tracked by time-lapse microscopy. At t0 min, only labeled cells exhibited a detectable fluorescence signal ( Figures 2 A and 2A ′). After 15 min, an apparent fluorescence signal was monitored from nonlabeled cells located in the vicinity of labeled ones ( Figures 2 B and 2B′). Remarkably, however, by t30 min almost all the nonlabeled cells displayed significant fluorescence while the fluorescence from labeled cells decreased ( Figures 2 C and 2C′ and Figure S3 Ba ). When unexposed regions of the growing cells were photographed at the latest time point, a similar fluorescence pattern was observed (data not shown). Consistently, labeled cells, located apart from nonlabeled ones, largely maintained their fluorescence signal ( Figure S3 A). Thus, same as GFP, calcein can be transferred from one cell to another; yet it appears to be delivered more rapidly, suggesting that the speed of transfer inversely correlates with the size of the traversed molecule.

(B) a: Average fluorescence intensity of nonlabeled B. subtilis cells as a function of their distance from calcein labeled B. subtilis cells at the beginning (0 min; light blue bars) and the end (30 min; dark blue bars) of a coincubation experiment as described in (A). No detectible signal was monitored when cells were located beyond 2 μm at 0 min. Average fluorescence signal is expressed in arbitrary units (AU). Error bars represent standard deviation (SD) of the mean fluorescence signal calculated from at least 40 cells located at the indicated distance. SD for calcein experiments was in general higher than that of the GFP experiments because of variation in labeling efficiency between individual cells. Shown is a representative experiment out of three independent biological repeats. b: Average fluorescence intensity of S. aureus (MRSA) (gfp−) cells as a function of their distance from B. subtilis (gfp+) cells at the beginning (0 min; light blue bars) and the end (50 min; dark blue bars) of a coincubation experiment (see Extended Experimental Procedures ). No detectible signal was monitored when cells were located beyond 2 μm at 0 min. Average fluorescence signal is expressed in arbitrary units (AU). Error bars represent SD of the mean fluorescence signal calculated from at least 40 cells located at the indicated distance. Shown is a representative experiment out of three independent biological repeats. c: Average fluorescence intensity of E. coli (gfp−) cells as a function of their distance from B. subtilis (gfp+) cells at the beginning (0 min; light blue bars) and the end (50 min; dark blue bars) of a coincubation experiment (see Extended Experimental Procedures ). No detectible signal was monitored when cells were located beyond 2.5 μm at 0 min. Average fluorescence signal is expressed in arbitrary units (AU). Error bars represent SD of the mean fluorescence signal calculated from at least 40 cells located at the indicated distance. Shown is a representative experiment out of three independent biological repeats.

(A) Exponentially growing PY79 cells were labeled with calcein. Labeled cells were washed and incubated in a temperature controlled chamber at 37°C (see Extended Experimental Procedures ). Cells were visualized by time-lapse fluorescence microscopy and images of phase contrast (blue) and fluorescence (green) were collected. Select fluorescence (a and b) and corresponding overlay images (a′ and b′) are shown from the following time points: (a and a′) t0 min (b and b′) t30 min. No significant loss of fluorescence occurred over time. However, when labeled cells underwent cell division (red arrow) or came in contact with a nonlabeled cell in the field (yellow arrow) loss of fluorescence was observed. This decrease in fluorescence is expected since, unlike GFP, calcein is not being regenerated. The scale bar represents 1 μm.

Exponentially growing PY79 cells were labeled with calcein (see Extended Experimental Procedures ). Labeled cells were washed and mixed with nonlabeled cells, plated on an LB agarose pad, and incubated in a temperature controlled chamber at 37°C. Cells were visualized by time-lapse fluorescence microscopy and images of phase contrast (blue) and fluorescence (green) were collected at 5 min intervals. Select fluorescence (A–C) and corresponding overlay images (A′–C′) are shown from the following time points: (A and A′) t0 min, (B and B′) t15 min, and (C and C′) t30 min. The scale bar represents 1 μm.

To further explore this phenomenon, time-lapse microscopy was carried out to follow the formation of the GFP gradient at a single-cell level. gfp+ and gfp− cells were mixed, applied to an agarose pad, and their growth and fluorescence were monitored. Immediately after mixing (t0 min), the fluorescence signal was confined to the gfp+ cells and no detectable fluorescence was seen in adjacent gfp− cells ( Figure 1 Ca). However, after 30 min, gfp− cells lying in proximity to gfp+ cells acquired a weak fluorescence signal ( Figure 1 Cb). The fluorescence intensity of gfp− cells increased over time in a manner inversely proportional to their distance from the gfp+ cells, i.e.: cells residing closer to the gfp+ cells acquired more fluorescence than distant ones ( Figure 1 D). Conversely, the fluorescence displayed by gfp+ cells decreased over time ( Figure 1 Cc), suggesting that they distribute their fluorescence among proximal cells. Observing a larger field highlights that as time progresses, gfp− cells not directly contacting gfp+ cells, also gained a fluorescence signal ( Figure S1 A available online). To rule out the possibility that these observations are a consequence of multiple fluorescence exposures, we imaged unexposed regions of the growing cells at the final time point, and a similar fluorescence pattern was detected ( Figure S1 B). Further, when gfp+ and gfp− cells were residing apart from one other, neither the gfp− cells gained nor the gfp+ cells lost fluorescence ( Figure S2 ). The contact-dependent nature of the fluorescence gradient excludes the possibility that the signal derives from cell migration and corroborates that cytoplasmic GFP molecules (27 kDa) can be transferred from one cell to another in a temporal and spatial manner. Of note, we cannot exclude the possibility that to some extent gfp transcripts are also being traded among the cells.

Exponentially growing PY79 (gfp−) and SB444 (gfp+) cells were mixed, plated on an LB agarose pad, and incubated in a temperature controlled chamber at 37°C. A region whereby gfp+ and gfp− cells were distant from each other was selected. Cells were visualized by time-lapse microscopy and images of phase contrast (blue) and fluorescence (green) were collected at 10 min intervals. Select overlay images are shown from the following time points: (A) t0 min (B) t20 min (C) t50 min (D) t70 min (E) t90 min and (F) t110 min. When gfp+ and gfp− cells are far apart, no fluorescence is detected from gfp− cells in all tested time points (dashed rectangular). However, a weak signal is monitored from gfp− cells residing in vicinity of gfp+ cells at the latest time point (indicated by arrows). The scale bar represents 2 μm.

(B) Exponentially growing cells of strains PY79 (gfp−) and SB444 (gfp+) were mixed, plated on an LB agarose pad, and incubated in a temperature controlled chamber at 37°C. Cells were photographed once, 60 min after the start of incubation. Left: An overlay image of phase contrast (blue) and fluorescence from GFP (green). Right: Fluorescence from GFP. The scale bar represents 2 μm.

(A) Exponentially growing cells of strains PY79 (gfp−) and SB444 (gfp+) were mixed, plated on an LB agarose pad, and incubated in a temperature controlled chamber at 37°C. Cells were visualized by time-lapse microscopy and images of phase contrast (blue) and fluorescence (green) were collected at 10 min intervals. Select overlay images are shown from the following time points: (a) t0 min (b) t30 min (c) t60 min and (d) t80 min. Paired arrows in (b) indicate locations where transfer of fluorescent molecules between neighboring cells is initiated. The level of fluorescence emanating from gfp− cells increases over time in a manner dependent on distance from gfp+ cells. Enlarged images of a selected area are shown in Figure 1 C. The scale bar represents 2 μm.

Given the complex intercellular communication required within natural bacterial communities, we reasoned that bacterial cells grown on a solid surface can physically interact in order to establish an effective route for exchange of molecular information. Initially, we examined whether adjacent cells exchange cytoplasmic GFP molecules. Bacillus subtilis cells (SB444) harboring a chromosomally encoded gfp reporter gene (gfp+) were spotted on solid medium alongside B. subtilis cells (PY79) lacking gfp (gfp−). Cells were allowed to grow for 15 hr and then visualized by fluorescence microscopy ( Figure 1 A ). Remarkably, a green fluorescence gradient was observed to emanate from the gfp+ cells toward the gfp− cells, covering a distance of approximately 40 μm ( Figures 1 Ab and 1B). Superimposing the green fluorescence and the phase contrast image, which demarcates the cells boundary, demonstrated that this fluorescence gradient was associated exclusively with the presence of cells ( Figure 1 Ac). The observed cell-associated gradient of the GFP signal concords with our premise that cytoplasmic molecular exchange occurs between neighboring cells. However, it remained possible that the gradient was due to migrating gfp+ cells.

(D) Average fluorescence intensity of the gfp− cells as a function of their distance from the gfp+ cells at t0 min (light blue bars) and at t60 min (dark blue bars) of the coincubation experiment as described in (C) (see Extended Experimental Procedures ). No detectible signal was measured when cells were located beyond 1μm at t0 min. Average fluorescence signal is expressed in arbitrary units (AU). Error bars represent standard deviation (SD) of the mean fluorescence signal calculated from at least 40 cells located at the indicated distance. Shown is a representative experiment out of three independent biological repeats.

(C) Exponentially growing PY79 (gfp−) and SB444 (gfp+) cells were mixed, plated on an LB agarose pad, and incubated in a temperature controlled chamber at 37°C. Cells were visualized by time-lapse fluorescence microscopy and phase contrast (blue) and fluorescence (green) images collected at 10 min intervals. Select overlay images are shown from the following time points: (a) t0 min, (b) t30 min, and (c) t60 min. Each pair of colored arrows (red and yellow) indicates different locations where transfer of fluorescent molecules between neighboring cells is increasing over time. Larger fields of the same region are shown in Figure S1 . The scale bar represents 1 μm.

(B) Average fluorescence intensity of the gfp− population (as indicated in Aa) as a function of the distance from the gfp+ population. The gfp− region was divided into identical sub-regions and the average fluorescence signal was defined in arbitrary units (AU).

(A) PY79 (gfp−) and SB444 (gfp+) cells were grown side by side on an LB agar plate at 37°C and visualized by fluorescence microscopy 15 hr after plating, when small colonies were visible. The dashed line indicates the border between the two populations. (a) Phase contrast image (blue). (b) GFP fluorescence image (green). (c) Overlay of phase and GFP fluorescence images. The scale bar represents 10 μm.

Discussion

We revealed the existence of a previously unidentified form of bacterial communication that facilitates the exchange of cytoplasmic constituents between adjacent cells via intercellular connecting tubes. Utilizing microscopy and genetic assays, we show that small cytoplasmic molecules and proteins can be traded between cells grown in proximity, thereby generating transient, nonheritable traits. Moreover, we demonstrate that nonconjugative plasmids can be transferred from one cell to another, resulting in transmission of hereditary features to recipient cells. Our data support that this type of communication is mediated by tubular projections that bridge neighboring cells and create a syncytium-like multicellular consortium. We propose that nanotube-mediated cytoplasmic sharing represents a key form of intercellular bacterial communication in nature, providing an efficient path for trading intracellular molecules between species. This attribute allows the emergence of new phenotypes by multispecies bacterial communities, increasing their survival in fluctuating environments.

Bassler and Losick, 2006 Bassler B.L.

Losick R. Bacterially speaking. Lazazzera, 2001 Lazazzera B.A. The intracellular function of extracellular signaling peptides. Ng and Bassler, 2009 Ng W.L.

Bassler B.L. Bacterial quorum-sensing network architectures. To date, the best characterized form of intercellular bacterial communication is known to be mediated by extracellular signaling molecules (). This type of communication is, however, constrained by the ability of bacteria to secrete and/or recognize the signal, transduce the received information, and modulate gene expression correspondingly. In contrast, communicating by nanotubes enables bacteria a straightforward immediate transfer of information that can cross the inherent species barrier. Moreover, because tunnels presumably maintain inner-cellular physiological conditions, channeled molecules are potentially protected from degrading enzymes and harsh environmental conditions. Taken together, nanotube-mediated informational flow is potentially both continuous and efficient and, as we show here, can enable molecular transfer across long distances in bacteria grown on solid surfaces ( Figures 1 A and 1B).

Madigan et al., 2003 Madigan, M., Martinko, J., and Parker, J., eds. (2003). Brock Biology of Microorganisms, 10th edn (Upper Saddle River, NJ: Pearson Education). Rosenshine et al., 1989 Rosenshine I.

Tchelet R.

Mevarech M. The mechanism of DNA transfer in the mating system of an archaebacterium. Schleper et al., 1995 Schleper C.

Holz I.

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Zillig W. A multicopy plasmid of the extremely thermophilic archaeon Sulfolobus effects its transfer to recipients by mating. Intercellular nanotubes connecting bacterial cells have been observed during conjugation, which is induced by conjugative plasmids and occurs in a unidirectional fashion from donor to recipient (). In contrast, the plasmid transfer described here does not require any intrinsic plasmid elements, and a given cell can be either donor or recipient. A similar phenomenon has been previously observed in archaebacteria, where nonconjugative plasmids were shown to reciprocally traverse from one cell to another and cytoplasmic bridges were detected between cells (). Therefore, nanotube-mediated plasmid transfer in bacteria, though less efficient than classical conjugation, most likely represents a more ubiquitous form of horizontal gene transfer in nature, enabling universal interspecies plasmid exchange without the need for a dedicated mechanism.

Davis and Sowinski, 2008 Davis D.M.

Sowinski S. Membrane nanotubes: dynamic long-distance connections between animal cells. Many questions remain unanswered concerning how cargo is transported through nanotubes. We do not know whether the transport is active and requires energy or is passive and prompted by diffusion. It is possible that both mechanisms coexist and utilization depends on the delivered cargo. In eukaryotic cells, nanotubes are frequently associated with cytoskeletal and motor proteins, implying a role for active transport (). The directionality of the transport is also elusive and raises the following questions: is there a defined donor and recipient, and does directionality depend on the cell that initiates tube formation? It will be interesting to explore whether a gating mechanism exists to control traffic directionality.

Our discovery that diverse bacterial species can communicate with nanotubes has significant medical implications. As we have demonstrated, both hereditary and nonhereditary antibiotic resistance can be acquired from neighboring cells through nanotubes, a survival strategy that could be widespread in nature. This unhampered informational flow raises the concern that nanotubes allow commensal bacteria to nurture pathogenic bacteria. Conversely, pathogenic bacteria may transfer virulence features to commensal bacteria converting them into pathogens. In this view, gaining a better molecular understanding of nanotube formation could lead to the development of novel strategies to fight against pathogenic bacteria.