Leptothrix pellicle with a distinct geometric pattern at the air-liquid interface.
Leptothrix cholodnii SP-6 and OUMS1 cells produce pellicles at the air-liquid interface under a static liquid culture condition (
Fig. 1A and see Movie S1 in the supplemental material) (
29), which has been reported for several bacteria. However, unlike pellicles consisting of a dense cell assembly, those of
L. cholodnii are porous structures with distinctive geometric patterns. To understand how such geometric patterns are formed, we separately inoculated SP-6 and OUMS1 into well dishes and imaged the process of their pellicle formation at high magnification at 1 min time intervals. The time-lapse image sequences indicate that tiny clusters of woven cell filaments sequentially gather at the outer edges of larger filament aggregates for both strains (
Fig. 1B and Movie S2). These observations lead us to conclude that the geometric patterning at the air-liquid interface is due to the random accumulation of tiny clusters of short cell filaments, a conserved feature for at least the
L. cholodnii strains.
The pellicle formation process in the dish culture is difficult to be viewed vertically from above because the surface-covering pellicles disturb focus stability, making it hard to observe the behavior of the cells and how the cells or cell filaments colonize underneath the liquid surface. For analysis of cellular activity such as the movement of small cell aggregates in the liquid phase, we devised equipment to image the cell culture horizontally as illustrated in Fig. S1. We separately inoculated SP-6 and OUMS1 cells into cuvettes and tracked the process of aggregate formation at 1 min intervals. The image sequences reveal that the cells of both strains predominantly accumulate at the air-liquid interface and produce a thin and rough layer within ~22 h (
Fig. 1C and Movie S3), which gradually gets thicker over time. However, we hardly observed small cell clusters in the liquid phases during pellicle formation (
Fig. 1C and Movie S3). Indeed, magnified images show that small aggregates of pellicles develop due to the trapping of planktonic cells by cell filaments (
Fig. 1D and Movie S4, left), indicating that single motile planktonic cells but not small aggregates may move toward the air-liquid interface. Next, we examined the requirement of flagella motility in pellicle formation.
Role of a polar flagellum-driven cell motility.
Leptothrix cells are known to have a single monopolar flagellum, which enables the planktonic cells to explore favorable sites (
19,
20). By constructing a deletion mutant of flagellar components, we investigated the role of the monopolar flagellum in the formation of floating pellicles. To construct the flagellum mutant, we prepared a spontaneous rifampin-resistant (Rif
r) mutant originating from SP-6 (the wild type [WT*], used for phenotype comparison to the disruptant) and its disruptant, in which
Lcho_2735 and
Lcho_2734, which encode putative components FlgA and FlgB, respectively, were deleted (Δ
flgAB cells). Since we removed the promoter region of the
flg operons encoding putative proteins FlgA through FlgM (
Lcho_2724 through
Lcho_2736), we expect expression of these proteins to be abrogated.
We confirmed the replacement of the flgA and flgB genes with the kanamycin resistance gene (Kmr) in candidate clones by colony PCR (Fig. S2A). Representative atmospheric scanning electron microscopy (ASEM) images revealed that the ΔflgAB cell loses its polar flagellum (Fig. S2B). Subsequently, we found that the ΔflgAB cells lose their cell motility driven by the flagellum. Movie S5 shows representative swimming planktonic WT* cells. In contrast, we did not observe swimming ΔflgAB cells, suggesting that the replacement of these genes with Kmr abrogates the flagellar function.
In shaking cultures, Δ
flgAB cells produced aggregates of cell filaments covered by a sheath, like the WT* (Fig. S2C and E) (
34). Contrary to WT*, Δ
flgAB cells failed to form porous pellicles in a static dish or cuvette cultures (
Fig. 2A and
B, Fig. S2D, and Movie S6). Using high-magnification time-lapse images, we found WT* aggregates and swimming planktonic cells at the air-liquid interface (
Fig. 2C, left). However, we observed several long cell filaments but no planktonic cells in the Δ
flgAB cell culture (
Fig. 2C, right). From these observations, we conclude that like
Bacillus (
16), the flagellum-driven motility of WT* assists planktonic cells to move toward the air-liquid interface and eventually contributes to expansion of the porous pellicles.
Dependency on sheath structure.
The
Leptothrix sheath is known to consist of nanofibrils and to have multiple functions, such as enabling surface attachment of planktonic cells to solid substrates, holding divided cells in line, mediating interactions between other cell filaments, and facilitating the encrustation of iron oxide-rich particles (
22–24). Furthermore, the nanofibrils elongating further outwards from the sheath structure efficiently expand its superficial area (
28), which might enable cell filaments to capture swimming planktonic cells as well as to facilitate assembly by the sequential docking with other small cell-filament clusters.
To explore whether nanofibril secretion is critical for the distinctive patterning of the net-like pellicle, we constructed a disruptant lacking the
Lcho_3510 gene (hereafter
lthA), encoding a putative exopolysaccharide biosynthesis polyprenyl glycosylphosphotransferase whose homolog,
sthA, is known to be responsible for the construction of the sheath structure in
Sphaerotilus natans (
33). After acquiring candidate clones of
lthA deletion mutant cells (Δ
lthA cells) using colony PCR, we confirmed the proper replacement of the gene (Fig. S3A). We further confirmed the absence of a sheath structure around the periphery of the Δ
lthA cell by atmospheric scanning electron microscopy (ASEM) (Fig. S3B) and fluorescent imaging of nanofibrils stained by Alexa Fluor 594 C5-maleimide (Alexa594-SH) (
Fig. 3A).
In a shaking liquid culture containing MSVP medium, the WT* strain generates fibrous filament aggregates. In contrast, ΔlthA cells are uniformly distributed in the entire medium, making it cloudy (Fig. S3C). Time-lapse imaging in a static dish culture reveals that the ΔlthA strain forms chains containing 4 or 5 cells at the bottom surface, which break into 1 or 2 cells within ~2 h (Fig. S3D, right). However, for the WT* strain, we observed very little fragmentation of chained cells (Fig. S3D, left). These observations indicate that the ΔlthA cells are not able to form sheathed cell filaments due to a failure of nanofibril secretion regardless of liquid culture condition.
In static dishes and cuvette cultures, unlike Δ
flgAB cells, Δ
lthA cells produce pellicles. Since planktonic Δ
lthA cells maintain their flagellum-driven motility, this could enable them to move up to the interface (Movie S5, right). Notably, Δ
lthA cells congregate to form a dense and relatively thick pellicle (
Fig. 3B and
C,
t =
40 h; Movie S7), which consists of cell assemblies that are structurally different from the net-like pellicle formed by WT* cells. Contrary to WT*, Δ
lthA pellicles release their resident cells into the liquid phase (
Fig. 3C,
t =
45 h; Movie S7). We use atomic force microscopy (AFM) to image dried samples from WT* and Δ
lthA pellicles. Confirming our expectations, we found that WT* cells are aligned in filaments, whereas the Δ
lthA pellicle cells, which lack nanofibrils, have aggregated (
Fig. 3D). This difference in aggregation morphology between WT* and Δ
lthA cells is likely due to differences in cell surface properties.
To estimate the cell density at the air-liquid interface during pellicle formation, we analyzed the interface area coverage and thickness of WT* and Δ
lthA pellicles from image sequences (Movie S7). In dish culture, the percentage of coverage of WT* pellicles plateaus at ~55% after ~30 h. In contrast, the percentage of coverage of Δ
lthA pellicles reaches 100% within the first 10 h, before collapsing into the liquid phase (
Fig. 4A and Fig. S4A and B). To facilitate measurement of pellicle thickness, we use cuvette cultures; these cultures will have the same qualitative features as dish culture, but will differ in terms of dynamics due to the differences in interface area. From time-lapse sequences, we find that the thickness of WT* pellicles reaches a constant value of 30 μm at ~25 h. In contrast, Δ
lthA pellicles demonstrate a long period during which thickness increases slowly, followed by a rapid rise and fall in thickness, which is due to pellicle collapse (
Fig. 4B and Fig. S4C and D). We find that Δ
lthA pellicles reach a maximum thickness of ~100 μm. From these values and from our surface coverage percentage, we estimate that at collapse, the Δ
lthA pellicles have ~6× more mass than WT* pellicles, assuming the same cell density for both strains. The rapid coverage at the air-liquid surface by Δ
lthA cells suggests that they have a higher cell surface hydrophobicity than WT* cells. Furthermore, due to the inability to form filaments, it appears that Δ
lthA cells can form much denser structures than WT* cells, which are confined to filamentous chains.
Adhesion at the air-liquid interface by pellicles can be mediated by surface-active agents (
17). However, the surface tension of the MSVP medium was nearly constant (≈71.5 mN/m) when WT* or Δ
lthA cells were cultured for 72 h (Fig. S5), suggesting that they were not secreting surface-active molecules. For
Bacillus subtilis, prior to pellicle formation, a transition from planktonic cells to nonmotile short cell chains has been reported (
16). Its EPS production assists the cell chain in adhesion and alignment that lead to the formation of a dense pellicle (
13). Our high-magnification images of the Δ
lthA pellicle reveal a loose packing of swimming planktonic cells at the air-liquid interface (Fig. S6), which over time transforms into nonmotile short cell chains (
t =
3 h) that eventually become the part of larger assemblies that have loose cell-cell interactions (
t =
5 h). This is distinct from SP-6 and WT* pellicles, which are composed of entangled cell filaments (
Fig. 1D). These observations indicate that sheathed cell chains are crucial for the formation of porous pellicles consisting of entangled cell filaments. The nanofibril-free, and thus sheathless, Δ
lthA cells form thick assemblies without the support of nanofibrils but are unable to form a net-like pellicle.
It has been reported that the glycoconjugates that compose the SP-6 nanofibrils are functionalized with dissociable SH- and NH
2-terminal groups that assist cell filaments in self-entanglement and adsorption of Fe-rich particles (
25,
27,
28). We investigated the effect of these terminal groups on the accumulation of tiny filamentous clusters at the air-liquid surface. We incubated SP-6 cells in MSVP containing Alexa594-SH and fluorescein-NH
2 reagents, which specifically block the binding of NH
2 and SH groups, respectively, by conjugation (
24,
28,
35) and tracked the formation of floating pellicles (Fig. S7). We observed no significant effect when blocking NH
2- and SH-terminal groups in the net-like SP-6 pellicles. We thus conclude that NH
2- and SH-terminal groups are not required for porous pellicle formation, although these play a primary functional role in interaction with iron oxides and nanofibrils, respectively (
27,
28).
Cell surface hydrophobicity and its impact on pellicle formation.
We have shown that sheathless Δ
lthA cells tend to self-aggregate compared to WT* cells (
Fig. 3D). Self-aggregation of bacteria is known to be primarily mediated by the recognition of surface components such as proteins and exopolysaccharides (
36). Moreover, cell surface hydrophobicity is also known to be an important factor for self-aggregation (
37). We therefore examined cell surface hydrophobicity using hydrophilic fluorescent sicastar-greenF microspheres (100 nm in diameter), which are designed to bind selectively to hydrophilic but not to hydrophobic substrates. We separately incubated sheathed WT* and sheathless Δ
lthA cells with 1 mg/mL of sicastar-greenF microspheres and then acquired fluorescence images. As we expected, a strong bonding of the sicastar-greenF to hydrophilic WT* cells resulted in an intense green fluorescence at the filament periphery (
Fig. 5A, upper; Fig. S8), while hydrophobic Δ
lthA cells showed a weak fluorescence signal, except at the edges of the cell (
Fig. 5A, lower). These results indicate that hydrophilic components are widely distributed on the cell surface of WT*. Furthermore, the orange fluorescence of Alexa594-SH, which blocks SH groups of nanofibrils in sheath structures, is perfectly matched with green fluorescence (
Fig. 5A), indicating that (i) nanofibrils, which form the sheath structures, are responsible for cell surface hydrophilicity, and (ii) SH groups in nanofibrils have no significant effect on their hydrophilicity. However, we could not determine whether the hydrophilicity of Δ
lthA cells is due to polar flagella that contain hydrophilic domains (
38).
To be fully assured of the cell surface hydrophobicity, we next examined the effect of nanofibrils and sheath structure on the cell surface hydrophobicity by measuring the two-phase cell-water-air contact angle by depositing a water droplet on a bacterial lawn (
39). We found that the average contact angle (θ) of the water droplet for Δ
lthA cells is 60% larger than that of the WT* cells (
Fig. 5B). This indicates that the sheath structure contributes to cell surface hydrophilicity, and thus sheathed WT* cells are more hydrophilic than sheath-free Δ
lthA cells.
Although several unknown factors may influence this result, cell hydrophobicity is meaningful for the pellicle formation process since it affects adsorption energy at the air-liquid interface. The adsorption energy (
E) of a bacterium adsorbed at an interface can be estimated from the equation
, where γ is the surface tension at the air-liquid interface,
S is the bacterium surface area, and θ is the air-liquid contact angle of cells. Here, the negative sign indicates energy is required for a cell to escape the interface (
40–42). We measured γ = 71.5 mN/m (Fig. S5) and
S = 6 μm
2 (
20). Evidently, for the same air-liquid interface, the adsorption energy of a bacterium mainly depends on its contact angle. Using the measured average value of the contact angle and the adsorption energy equation, we estimate that Δ
lthA cells are adsorbed ~5 times more strongly than the WT* cells. This stronger adsorption of Δ
lthA cells appears to reduce both swimming motility and Brownian motion, as is visible in Movie S4, which shows a rotational motion of planktonic Δ
lthA cells, while aggregates of Δ
lthA cells do not show any motion. Furthermore, due to the larger value of Δ
lthA cell hydrophobicity, these cells will deform the air-liquid interface more than WT* cells (
43–45). Over time, due to the “Cheerios effect,” in which interfacial particles attract each other due to surface tension, Δ
lthA cells may attract each other and form a dense pellicle (
46), as is visible in Fig. S5. The reason for the rotational motion of planktonic Δ
lthA cells (Movie S4, right) is not clear, but we speculate it may be due to a countercoupling of flagellar motile energy and interface adsorption energy. In contrast, weaker adsorption of hydrophilic planktonic WT* cells and their flagellar motility allows them to move and explore the interface, as can be seen in Movie S4 (left). While in motion, they randomly stick to the surface by adhesive nanofibrils and initially develop small aggregates that further enlarge as small aggregates stick together and develop net-like pellicles.
Conclusion and outlook.
In this study, we investigate the mechanism of pellicle formation by the filamentous bacterium Leptothrix cholodnii SP-6 at the air-liquid interface. Our findings show that the flagellar motility of WT* cells is primarily responsible for transporting cells from bulk liquid to the interface and that cell surface hydrophilicity in part controls the pellicle density and shape. We find ΔlthA cells that were unable to produce sheath structures are 60% more hydrophobic than the sheathed WT* cells, showing that the presence of the sheath structure increases cell hydrophilicity. The estimation of air-aqueous interface adsorption energy shows hydrophobic ΔlthA cells are adsorbed ~5x more strongly than WT* cells. This larger adhesion energy inhibits their movement away from the interface, where they aggregate into the dense pellicle. In contrast, weaker adsorption of WT* cells enables them to swim to, explore, and depart from the interface. During this movement, the planktonic WT* cells stick to one another due to their filamentous structures and form small aggregates that gradually enlarge by the random docking of smaller aggregates, forming branched pellicles.
Bacterial pellicle formation is shown to be triggered by inducers such as iron for
Pseudomonas fluorescens, dimethyl sulfoxide or ethanol for
E. coli, and arabinose for
Vibrio fischeri (
47–49). Therefore, it might be possible that the pellicle formation of
L. cholodnii is induced by minerals such as iron oxides, but further studies are needed to identify the factors required for pellicle formation. In addition, since we have reported that sheath-forming bacteria
Leptothrix,
Sphaerotilus, and
Thiothrix excrete different types of glycoconjugates (
25,
50,
51), we plan to investigate if there are differences in sheath hydrophobicity in future studies.
The filamentous bacteria of the genera
Leptothrix and
Sphaerotilus are widely distributed in wastewater treatment plants (
19,
21). However, their proliferation and aggregation often clog water distribution systems, which reduces the efficiency of the treatment plants (
52). The findings of the present study provide a deeper understanding of biological and physiochemical factors that influence pellicle formation. We hope to contribute to finding an effective method to control their proliferation that may be employed in large-scale applications.