Skip to main content
eLife logoLink to eLife
. 2022 Oct 11;11:e71920. doi: 10.7554/eLife.71920

Novel multicellular prokaryote discovered next to an underground stream

Kouhei Mizuno 1,2,, Mais Maree 3, Toshihiko Nagamura 2, Akihiro Koga 2, Satoru Hirayama 4,5, Soichi Furukawa 4,, Kenji Tanaka 6, Kazuya Morikawa 7,
Editors: Raymond E Goldstein8, Detlef Weigel9
PMCID: PMC9555858  PMID: 36217817

Abstract

A diversity of prokaryotes currently exhibit multicellularity with different generation mechanisms in a variety of contexts of ecology on Earth. In the present study, we report a new type of multicellular bacterium, HS-3, isolated from an underground stream. HS-3 self-organizes its filamentous cells into a layer-structured colony with the properties of a nematic liquid crystal. After maturation, the colony starts to form a semi-closed sphere accommodating clusters of coccobacillus daughter cells and selectively releases them upon contact with water. This is the first report that shows that a liquid-crystal status of cells can support the prokaryotic multicellular behavior. Importantly, the observed behavior of HS-3 suggests that the recurrent intermittent exposure of colonies to water flow in the cave might have been the ecological context that cultivated the evolutionary transition from unicellular to multicellular life. This is the new extant model that underpins theories regarding a role of ecological context in the emergence of multicellularity.

Research organism: Other

Introduction

The emergence of multicellularity is one of the mysteries of life on Earth, and has been attracting scientists in a broad range of fields including bioengineering on artificial construction of cellular behaviors (Basu et al., 2005) and medical research on cancer progression (Trigos et al., 2017). The oldest reliable fossil record of multicellular life is an ~3.42-billion-year-old putative filamentous microfossil that inhabited a paleo-subseafloor hydrothermal vein system (Cavalazzi et al., 2021). Since then, the fossil records show the emergence of dominating multicellular cyanobacteria on this planet (Amard and Bertrand-Sarfati, 1997; Golubic and Seong-Joo, 2004), which were identified as early as 2.32 billion years ago and are assumed to be responsible for the rapid accumulation of oxygen levels, known as the ‘Great Oxygenation Event’ (Schirrmeister et al., 2011). It seems that multicellularity emerged several times independently at different species lineages irrespective of a eukaryotic or prokaryotic phenotype (Brunet and King, 2017). Multicellularity in prokaryotes would not directly link to the origin of animals, but investigating it gives us fundamental models to consider the mechanistic or evolutionary steps behind the leap to multicellularity. Currently, we see a diversity of prokaryotes that exhibit multicellularity with distinct formation mechanisms in different contexts of ecology on Earth (Claessen et al., 2014; Lyons and Kolter, 2015; Nadell and Bassler, 2011; Vlamakis et al., 2013; Zhou et al., 2013; Figure 1).

Figure 1. Prokaryotic multicellularity.

Figure 1.

‘Clonal multicellularity’, ‘aggregative multicellularity’ (Brunet and King, 2017), and ‘multicellular-like behavior’ (Malik and Rosenberg, 2009) describe modes of multicellularity that arose through distinguished hypothetical conditions.

Actinomyces and cyanobacteria are well-studied prokaryotes that exhibit multicellular forms (Claessen et al., 2014). Their reproducible multicellular structures achieve a high fitness for the whole population even though they are not beneficial for partial individuals. Further elaborate organization can be seen in large-sized genome bacteria, such as Myxobacteria (9–16 Mbp genome) and Streptomyces (7–12 Mbp) (Claessen et al., 2014). Some fitness advantages acquired by multicellularity include avoiding predation (Bernardes et al., 2021; Boraas et al., 1998; Herron et al., 2019), cooperative defense (Shapiro, 1998), dispersal (Smith et al., 2014), labor division (Brunet and King, 2017; Shapiro, 1998; Koschwanez et al., 2011), resource competition (Herron et al., 2019), and creation of a stable internal environment (Brunet and King, 2017).

The terms ‘aggregative multicellularity’ or ‘clonal multicellularity’ are used to describe the different bottom-up processes resulting in multicellular structures (Figure 1). An example of aggregative multicellularity can be seen in myxobacteria, where solitary cells come together upon stress conditions to organize a fruiting body (Bonner, 1998). On the other hand, clonal multicellularity occurs through incomplete cell division, such as in actinomyces and cyanobacteria, and may be the process by which the first eukaryotic ancestor of all animals evolved (Brunet and King, 2017). The co-option of preexisting cellular traits is an important concept in these bottom-up processes; for example, cell aggregation in response to stress, or a sticky cellular trait might have been used to architect a multicellular status (Brunet and King, 2017; Malik and Rosenberg, 2009).

Once a multicellular organism has emerged, the multicellular trait needs to be internalized as genetic information and fixed in the population for which Darwinian natural selection, which occurs over a longer timescale encompassing multiple cell lifetimes, must take place (Black et al., 2020). This is a key problem of the bottom-up perspective of multicellularity (Libby and Ratcliff, 2014). However, the constraints that allow for the selection of a multicellular status are as yet unknown. One suggested explanation of the fixation of multicellularity is the involvement of certain environmental factors that can give an extrinsic selective pressure to the evolving population. This concept is termed ‘Ecological Scaffolding’ (Black et al., 2020) and has been experimentally substantiated in a model experiment using Pseudomonas (Hammerschmidt et al., 2014). The study regarded a biofilm formed at the air–liquid interface of a glass bottle as a population at a certain generation, and counted repeated dynamism of synthesis and collapse of the biofilm as generations of the population. This concept suggests that Darwinian natural selection is possible even for unicellular organisms through support by external environmental constraints before multicellular traits are fixed in a population.

The cave microbiome was previously considered to simply be a subset of the surface microbes (Laiz et al., 1999), but it has recently been shown that the overlap of operational taxonomic units with microbes from the surface is only 11–16% (Lavoie et al., 2017; Ortiz et al., 2013). Furthermore, the microbiomes of different cave environments differ, as has been revealed in caves composed of limestone (Barton and Northup, 2007; Ortiz et al., 2013, Rangseekaew and Pathom-Aree, 2019), lava (Lavoie et al., 2017), ice (Itcus et al., 2018), quartz (Sauro et al., 2018), or sulfur (Macalady et al., 2008). Namely, each cave has a distinctive microbiome that has adapted to that environment.

Here, we report a new cave bacterium that can develop a multicellular-like architecture, possibly through the influence of an external environmental constraint, from a simple, but ordered, cell cluster. The bacterium was isolated from the surface of a cave wall above an underground river in a limestone cave system that is intermittently submerged. The most remarkable aspect of this bacterium’s life-cycle is the existence of well-regulated development (dimorphism) between the first growth stage (filamentous cells that show liquid-crystal-like self-organization on a solid surface) and the second stage (coccobacillus cells that can disperse in flowing water). There is no report that the properties of a nematic liquid crystal were co-opted toward prokaryotic multicellularity. The second stage suggests the involvement of recurrent water flows in the establishment of the multicellularity in this species. We discuss what the discovery of HS-3 implicates in the context of our present understanding and hypothesis about the emergence of multicellularity.

Results

Jeongeupia sacculi sp. nov. HS-3, a new bacterial species isolated from a karst cave

We previously collected oligotrophic bacteria from a variety of environments and studied their physiology (Ilham et al., 2014; Mizuno et al., 2017; Mizuno et al., 2010). In 2008, strain HS-3 was isolated from water dripping on a limestone cave wall in the Hirao karst plateau (Fukuyama et al., 2004) in northern Kyushu Island, Japan (Figure 2—figure supplement 1). The sampling site, which was located slightly above the water surface of an underground river, was intermittently submerged due to an increase in runoff after rainfall (Figure 2A, Figure 2—figure supplement 1). After plating on agar, colonies appeared transparent and iridescent (Figure 2B). HS-3 is a Gram-negative obligate aerobe with a single polar flagellum (Figure 2F, inset). Genetically, it belongs to a group in the family Neisseriaceae that is mainly comprised of environmental bacteria (Figure 2—figure supplement 2). It has a 3.4 Mbp genome and a plasmid (2.0 kbp). Based on phenotypic comparisons with other closely related species (Supplementary file 1), HS-3 is considered to be a novel species, and we name this species as Jeongeupia sacculi sp. nov. HS-3. Its optimal growth temperature (24°C) and pH (8.0–9.0) are consistent with conditions in the cave. Another species in the same group that is adapted to life in oligotrophic environments is Deefgea rivuli, which has been isolated from tufa, a form of limestone (Stackebrandt et al., 2007; Figure 2—figure supplement 2A). However, no reports of distinctive colony morphology or cell differentiation have been reported in this group to date.

Figure 2. Liquid-crystal appearance and microstructure of a colony of Jeongeupia sacculi sp.nov.

HS-3, discovered in a limestone cave. (A) River in a cave showing the sampling site on the cave wall (inset). (B) Transparent and iridescent appearance of colonies. The edge of a colony growing on agar. Differential interference contrast (DIC) (C) and scanning electron microscopic (SEM) (D, E) images. Bulges of cells on the colony surface (E). (F) Cell length in liquid and solid cultures. The liquid cultures were monitored at different times. The solid cultures were inoculated at different colony densities (CFU/plate) and cultured for 2 days. The value given for each sample is the mean obtained from 100 cells. Inset: transmission electron microscopic (TEM) image of a liquid-cultured cell. (G) Typical cell morphology from a colony cultured on agar and observed by green fluorescent protein (GFP) fluorescence. Arrows: bulges of cells.

Figure 2.

Figure 2—figure supplement 1. The isolation site of strain HS-3.

Figure 2—figure supplement 1.

Images of the site where the bacterium was collected in a limestone cave: (A) Map of Japan. Arrowhead: Kyushu Island. (Inset) Arrowhead: Hirao karst plateau; (B) Aerial photograph of the sampling site in the Hirao karst plateau, the whole area of which measures 6.5 km from north to south and 2.5 km from east to west, and has an altitude of between 300 and 700 m. Arrowhead: Seiryu limestone cave. The aerial image was obtained from Geospatial Information Authority of Japan; (C) Photograph of Hirao karst plateau; (D) Cave wall (sampling site); (E) Entrance viewed from inside the cave; (F) Entrance from outside the cave; (G–I) River flowing in the cave. The air temperature inside the cave was around 15°C. The pH of the water on the wall was 8.5.
Figure 2—figure supplement 2. Phylogenetic analysis and genome information of HS-3.

Figure 2—figure supplement 2.

(A) Phylogenetic tree based on 16S rRNA gene sequence. The tree represents a cluster in the Neisseriaceae family (shadowed area) and some relatives isolated from similar oligotrophic environments such as tufa, another form of limestone, as indicated by blue arrows. The closest relatives of strain HS-3T are Jeongeupia naejangsanensis KCTC 66233T (FJ669217) and Jeongeupia chitinilytica JchiT (NR_109420) with similarity values of 98.2% and 97.8%, respectively. (B) Diagrams of the genome and the plasmid. The plasmid seems to be a low copy number one for which it has not been experimentally detected by a commercial extraction kit.

HS-3 forms colonies with a liquid-crystal texture

Bacterial cells capable of growth on agar medium typically proliferate in a disordered state, yielding colonies with an opaque texture. HS-3 formed transparent colonies with an iridescent hue (Figure 2B). Specifically, the colonies assumed an anisotropic optical structure (Figure 3E), suggesting that the cells were orientated in a manner similar to liquid-crystal molecules. As shown in Figure 2C–E, the cells in the colonies formed vortex-shaped structures composed of closely arrayed filamentous cells with bulges on the surface. Transforming these cells with the green fluorescent protein (GFP) gene revealed that these structures were composed of cells rather than some extracellular matrix or artifacts derived from electron microscopy (Figure 2G). On a solid medium, the length of the cells was correlated with the colony density, and lower densities were associated with a wider range of cell length (Figure 2F). These findings suggest that cell elongation in this strain is a regulated process that occurs in response to environmental cues. Cell elongation was not observed in liquid cultures.

Figure 3. Colony morphogenesis and liquid-crystal-like ordering.

(A) A microcolony showing a single layer of cells. Differential interference contrast (DIC) (top left) and green fluorescent protein (GFP) (top right) images and an enlarged image (bottom) of the area indicated by the white box. (B) A young colony (larger than A) with a single layer of cells at the edge. Enlarged image of the area indicated by the white box (right). (C) Anisotropic pattern of the bottom cell layer of a single colony. (D) The color mapping to show the orientation of cells. The anisotropic pattern is a typical characteristics in a nematic phase (Doostmohammadi et al., 2018). (E) Anisotropic optical texture of colonies of HS-3 (left) and Jeongeupia naejangsanensis (right), the close relative strain. DIC images (left) and diagonally illuminated images (right). Arrows indicate the direction of light illumination. (F) Time-lapse images of liquid-crystal-like pattern formation on the edge of a growing colony. Time (hr) after chase is indicated. The original video is available as Video 1.

Figure 3.

Figure 3—figure supplement 1. Emergence of analogous nematic patterns on the colony edge.

Figure 3—figure supplement 1.

(A) The emergence of analogous nematic patterns at the colony edge. Two boxed areas are shown as examples on the right. The areas are chronologically aligned and show two types of topological defects characterized by two-dimensional nematic liquid crystals. (B) Time course images of the enlarged area indicated by a box on the colony edge. A key feature underlying the development of the unique cellular architecture is related to the cells adopting the two-dimensional, nematic liquid-crystal orientation (Doostmohammadi et al., 2018), in which cells are aligned tightly together to form the layered structure. Other bacteria, such as E. coli and B. subtilis, can produce nematic features, but only for short periods when the cells are allowed to grow two dimensionally (Dell’Arciprete et al., 2018; Yaman et al., 2019). In these species, collisions between the expanding layer of cells results in edge instability and buckling of the two-dimensional nematic order. Conversely, HS-3 filamentous cells have the remarkable ability to sustain an ordered structure.

A time course analysis of the growing colonies showed that the cells proliferate as coccobacilli for the first 10 hr (data not shown). Then, as cell elongation was initiated from the edge of the colony, the cells gradually assumed topological characteristics similar to the two-dimensional nematic pattern described in liquid-crystal theory (Doostmohammadi et al., 2018; Duclos et al., 2017), with comet-like (+1/2) and trefoil-like (−1/2) topological defects being visible (Figure 3—figure supplement 1). Generally, the microcolonies initially spread as a single layer. As the colony expands, bulges are produced especially, but not specifically, around the rapidly growing colony edge (Figure 2E). We consider that the morphological change occurs to sustain the ordered state by relieving internal physical pressure. The disturbed single layer at the center then serves as the focal point for the expansion of additional layers (Figure 3A, differential interference contrast). The bulges are also generated around the colony center (data not shown). As the multilayered colony continues to grow, the expanding edge swallows the adjacent region and the internal filamentous cells buckle, generating domains with a vortex structure (Figure 3F, Video 1). Importantly, the filamentous cells appear to sustain the well-aligned structure, suggesting that there is tight adhesion among the filamentous cells (Figure 2E). An anisotropic pattern that emerged on the bottom layer of the colony was sustained throughout the period of colony growth (Figure 3C, D). Furthermore, the optical characteristics of the multilayer colonies indicate that the ordered structures are sustained (Figure 3E).

Video 1. The edge of a colony.

Download video file (13.6MB, mp4)

Internal proliferation to germ-like phase

Colonies stopped growing at 2 days after inoculation and no further changes were observed for the following 2–3 days. However, internal proliferation then started and the colony began to swell three dimensionally (Video 2). Internally proliferating cells formed optically distinct regions within a transparent colony, suggesting the loss of the ordered-layer structure. Nile red, a lipid-staining fluorescent dye, was used to selectively stain the membranes of the elongating cells in a colony (Strahl et al., 2014), and the internal coccobacillus cells were recognized as nonstained areas in three-dimensional cross-sectional images (Figure 4A), as well as in micrographs obtained by conventional fluorescence microscopy (Figure 4B). At the fifth day after inoculation, the internal cells were crowded out of the colony in our experimental setting (Figure 4B, right panels). The occurrence of this event in a single colony triggered a chain reaction of this phenomenon in adjacent colonies (Figure 4C), indicating some form of control.

Figure 4. Internally growing cells and crowding out.

(A) Three-dimensional image of a 4-day-old colony reconstructed from 100 Z-axis slices with a total height of 168 µm. The whole Nile-red image (left) and its cross-sections (right). (B) Five-day-old, mature colonies showing internal growth at the center of the colony. Two right panels show the ‘crowding out’. (C) Colony-crowding influences adjacent colonies by inducing a ‘chain reaction’. The direction of the chain reaction and its front line are indicated by the blue arrow and the dotted line, respectively. More details of a crowding colony are shown in Video 2. (D) Time-lapse images of a mature colony in air (left panel) and after being submerged in water (middle and right panels). The arrow indicates the waterborne coccobacillus cells released from the colony. The colonies were grown for 5 days on agar before the time-lapse analysis. The time (min) after chase is shown. Bars = 100 µm. The original video is available as Video 3. (E) Illustration of the potential multicellular life-cycle of HS-3. (I) Coccobacillus cells attached to a new solid surface. (II) The cells proliferate to some extent, generating a liquid-crystal texture. (III) The center area becomes raised due to clustering of internal cells, leading to the germ-like phase. (IV) A waterflow releases coccobacilli. (V) In the absence of a waterflow, outward growth of the internal coccobacillus cells results in ‘crowding out’.

Figure 4.

Figure 4—figure supplement 1. Reversibility between filament and coccobacillus cells.

Figure 4—figure supplement 1.

(A) HS-3 crowds out coccobacillus cells from mature colonies. Streaking of the daughter coccobacillus cells on a new plate showed that the coccobacilli could reproduce the original colony morphology with filamentous cells. Phase contrast images (top left) and a differential interference contrast (DIC) image (top right), and scanning electron microscopic (SEM) images (second row). (B) The filamentous cells were selected from a single 2-day-old colony, suspended in phosphate-buffered saline (PBS), and inoculated on a new agar plate. The mature filament cells that survived physical damage during inoculation grew by dividing on agar. The microscopic observation was performed without a cover slip. The time after chase is shown. Bars = 50 µm.

Video 2. Emergence of coccobacillus cells.

Download video file (7.9MB, mp4)

The sampling site of HS-3 was a moist cave wall above an underground river (Figure 2A). Since this site is also intermittently submerged by water due to increased runoff caused by rainfall, we examined the response of colonies on an agar plate to being submerged by water. Remarkably, the coccobacilli were released into the water column, leaving the architecture produced by the filamentous cells behind (Figure 4D, Video 3). Even after the complete release of the coccobacillus from the center of the colony, the bottom cell layer of the aligned filamentous cells adhered tightly to each other (Figure 4D, right side). These observations suggest that the filamentous cells act to support the embedded coccobacilli clusters. We also examined whether the released ‘daughter’ cells were able to reproduce the original colony morphology. The results showed that the spawned coccobacillus cells could indeed reproduce the original filamentous colony morphology (Figure 4—figure supplement 1A). In addition, when filamentous cells obtained from a mature colony were inoculated on agar (Figure 4—figure supplement 1B) the majority of cells failed to regrow, potentially due to cell damage during recovery. However, a fraction elongated and then divided to generate short rods that each formed a microcolony (Figure 4—figure supplement 1B). Thus, the filamentous cells were not irreversibly destined to be the equivalent of ‘somatic’ cells. These results suggest that the different morphological changes observed in the life-cycle of HS-3 are reversible (Figure 4E).

Video 3. Waterborne coccobacillus cells from a colony.

Download video file (13MB, mp4)

Discussion

There has been a wide variety of work regarding the emergence of multicellularity on extant multicellular organisms such as volvocine green algae and holozoans (e.g., choanoflagellates and ichthyosporeans) (Bonner, 1998; Brunet and King, 2017; King et al., 2008; Matt and Umen, 2016; Sebé-Pedrós et al., 2016). While it appears to have occurred both repeatedly and independently in different lineages, the generative process is enigmatic (Brunet and King, 2017; Richter et al., 2018; Shapiro, 1998); moreover, we cannot directly observe the transmission process of extant organisms, including HS-3. Nonetheless, in this study, we found unique characteristics of HS-3 that have some implications regarding the emergence of multicellularity.

The remarkable aspect of HS-3’s life-cycle is the regulated dimorphism between the first growth stage (filamentous cells showing liquid-crystal-like self-organization) and the second stage (coccobacillus cells). The first stage suggests that liquid-crystal-like self-organization is involved in the emergence of multicellularity, which has not been reported before. The liquid-crystal behavior of microscopic cell populations is recognized as the spontaneous formation of intrinsic two-dimensional patterns when motile, rod-shaped cells proliferate in thin layers at a certain density. Typical two-dimensional nematic patterns are characterized by asymmetric vortex structures and anisotropic domains adjacent to specific types of topological defects (Duclos et al., 2017; Majumdar et al., 2014; Sanchez et al., 2012); this was also observed for HS-3 colony on solid surface (Figure 3—figure supplement 1). The rise of the fluid dynamics subdiscipline in physics led to a number of theoretical and experimental approaches for cell behaviors in the last two decades, where researchers observed liquid-crystal orderings in cell suspension of neuronal cells, fibroblasts, melanocytes, osteoblasts, lipocytes, and several laboratory model bacteria such as Escherichia coli and Bacillus subtilis (Doostmohammadi et al., 2018). In ordinary bacteria as observed in E. coli growing on agar, the nematic features are only sustained for short periods (Dell’Arciprete et al., 2018; Yaman et al., 2019). In contrast, HS-3 is unique in its ability to maintain the two-dimensional sheet structure for a prolonged period, which we consider to be one of the prerequisites for HS-3 to establish its multicellular behavior.

This prolonged nematic status enables HS-3 to make its colony growth a two-phase life-cycle. Two distinct life-cycles are seen in extant single-celled bacteria such as spore formation in B. subtilis and asymmetric cell division of Caulobacter crescentus (Casadesús and Low, 2013; Dubnau and Losick, 2006). More generally, bacteria produce differentiated subpopulations using a variety of genetic (e.g., phase variation) or nongenetic (e.g., bistability) mechanisms (Casadesús and Low, 2013; Dubnau and Losick, 2006), by which they can achieve division of labor or hedge their bets in the event of unpredictable future conditions (Malik and Rosenberg, 2009). Although these dichotomic behaviors at population level underlie in multicellular life, bacteria still require some more process to acquire and establish distinctive multicellular structures. In the case of HS-3, we observed that internal growth of coccobacilli was attenuated until the colony of fibrous cells matured, and this regulation is likely a factor that enabled the establishment of multicellular-like structure.

Another important condition to support HS-3’s multicellular-like behavior appears to be the dynamic water environment in the cave, which we believe is the key to explain a potential evolutionary path toward HS-3’s multicellularity. The recurrent dynamism of water flow could have allowed each colony on the wall (i.e., a collective of genetically homogenous population) to undergo Darwinian natural selection. This assumption gives an attractive explanation as to how a genetically homogeneous single-cellular organism might have experienced natural selection through a benefit during the early stage of the population characteristics, and supports the ‘Ecological Scaffolding’ theory (Black et al., 2020).

In summary, the present study underpins the aforementioned theories and concepts about the transition from unicellular to multicellular life. HS-3 is an example of an extant prokaryote with a unique multicellular behavior that is based on liquid-crystal characteristics and timely regulated development, and potentially involved recurrent dynamism in its cave environment in the emergence of multicellularity.

Materials and methods

Strains and culture media

Strain HS-3 was isolated in 2008 from percolating water on the surface of a limestone cave wall in ‘Seryukutsu’ (Blue Dragon) cave in the Hirao karst plateau (33°46′00.5″ N 130°55′02.7″ E) in northern Fukuoka Prefecture, Japan (Figure 2—figure supplement 1). The sampling site on the limestone wall was situated close to the surface of an underground river. Water dripping from the wall was collected using a sterile plastic tube and the pH and temperature were measured on site. Samples were then stored at 4°C for several days until analysis. The water sample was diluted with phosphate-buffered saline (PBS, pH 7.0) to inoculate R2A (0.5 g yeast extract, 0.5 peptone, 0.5 casamino acids, 0.5 glucose, 0.3 K2HPO4, 0.05 MgSO4 per liter of deionized water) agar plates containing Nile red (Sigma-Aldrich, Missouri, USA) at 0.5 mg per liter. The HS-3 strain was originally isolated as a Nile red-positive strain, indicating that it was a lipid producer. The isolated HS-3 strain was precultured in R2A liquid medium in a test tube overnight at 24°C in a reciprocal shaker at 100 rpm. Then, the culture was diluted with PBS and inoculated on a plate containing R2A and 1.5% agar for taxonomic and microscopic analysis. The close relatives, Jeongeupia naejangsanensis KCTC 22633T and J. chitinilytica KCTC 23701T were purchased from the Korean Collection for Type Cultures, Taejon, Korea. Two closely related bacteria, Andreprevotia chitinilytica NBRC106431T and Silvimonas terrae NBRC100961T, were purchased from the National Bio-Resource Center, Kisarazu, Japan. Those strains were cultured in R2A medium at 24 or 30°C on agar, or in a reciprocal shaker at 100 rpm for liquid cultures.

Taxonomic identification of strain HS-3

The Gram reaction was performed using a Gram Staining Kit (Sigma). Culture media and conditions for taxonomic comparisons were essentially the same as those used for the reference strains, J. naejangsanensis KCTC 22633T (Yoon et al., 2010) and J. chitinilytica KCTC 23701T (Chen et al., 2013). Growth at various temperatures (4, 10, 16, 18, 20, 22, 24, 26, 28, 30, 32, 34, and 40°C) and pH (3.0–11.0 at intervals of 0.5) was performed in nutrient broth (Becton Dickinson, New Jersey, USA). The pH was adjusted by using sodium acetate/acetate, phosphate, and Na2CO3 buffers. Growth at different NaCl concentrations (0, 0.5, 1.0, 2.0, 3.0, 4.0, and 5.0%, wt/vol) was also performed in tryptic soy broth prepared according to the formula used for the Difco medium, except that NaCl was excluded. Growth under anaerobic conditions was performed using an anaerobic chamber with a gas exchange kit (Anaeropack; Mitsubishi Gas Chemical, Tokyo, Japan). Catalase and hydrolysis of casein, gelatin, starch, and Tween 20, 40, 60, and 80 were determined as described by Cowan and Steel, 1965. Additionally, assimilation of Tween 20, 40, 60, and 80 as sole carbon sources was tested in M9 medium (12.8 g Na2HPO4⋅7H2O, 3 g KH2PO4, 0.5 g NaCl, and 1 g NH4Cl per liter of deionized water containing 2 mM MgSO4 and 0.1 mM CaCl2). Susceptibility to antibiotics was tested by placing antibiotic-impregnated discs on agar plates seeded with the test strain. The antibiotics tested were kanamycin, gentamycin, erythromycin, fosfomycin, trimethoprim, novobiocin, streptomycin, chloramphenicol, ampicillin, and tetracycline. Utilization of various substrates and biochemical properties were tested by using API 20NE, API 50CH, API ZYM kits (bioMerieux, Marcy l'Etoile, France). Major types of quinones were determined at Techno Suruga, Shizuoka, Japan, using reverse-phase high-performance liquid chromatography (HPLC) (Komagata and Suzuki, 1988). Cellular fatty acid composition was also determined at Techno Suruga using the MIDI/Hewlett Packard Microbial Identification System (Sasser, 1990). The strain was deposited at the National Bioresource Center, Kisarazu, Japan as Jeongeupia sp. HS-3 NBRC 108274T (=KCTC 23748T).

Genomic DNA was extracted using a Qiagen kit (Qiagen, Hilden, Germany), and the 16S rRNA gene was amplified using the primer set of 27 f and 1525 r (Lane, 1996). The PCR product was sequenced (Applied Biosystems 3730xl DNA Analyzer, Applied Biosystems, USA), and the DNA G + C content was measured by Techno Suruga Inc using HPLC (Katayama-fujimura et al., 2014). A phylogenetic tree was constructed using the neighbor-joining method (Saitou and Nei, 1987) implemented in the Clustal W software package (Thompson et al., 1994) using sequences of closely related bacteria in GenBank that were selected by BLAST (Altschul et al., 1997).

Genomic DNA for genome sequencing was extracted using a NucleoSpin kit (Macherey-Nagel, Düren, Germany). Complete sequencing was performed at the Oral Microbiome Center, Takamatsu, Japan, using a combination of long-read sequencing performed using a Nanopore system (Oxford Nanopore Technologies, Oxford, UK) and short-read sequencing performed using a DNBSEQ system (MGI Tech, Shenzhen, China). The sequence data were analyzed using Lasergene software (DNASTAR) and annotations were performed by DFAST (https://dfast.nig.ac.jp/). The complete genome sequence and a plasmid sequence have been registered in the DDBJ database (https://www.ddbj.nig.ac.jp/) as AP024094 and AP024095, respectively.

GFP recombination

Competent cells of strain HS-3 were prepared using general laboratory protocols (Sambrook and Russel, 2001). Briefly, cells cultured in liquid R2A medium for 48 hr at 24°C were inoculated into a fresh R2A medium at a dilution of 20 times. Then, the cells were cultured until the OD600 nm reached 0.5. The cells were harvested by centrifugation and washed two to three times with sterile water and 10% glycerol, with a gradual increase in the concentration of the suspended cells. Finally, the cells were suspended in 10% glycerol, distributed to tubes, and stored at −80°C. The broad-host-range plasmid, pBBR1MCS2-pAmp-EGFP (Kovach et al., 1994) (Nova Lifetech, Hong Kong, China), was transformed into competent cells of HS-3 by electroporation under the following conditions. A mixture containing 150 ng of the plasmid and 100 µl of the cell suspension was placed in a cuvette with a 2.0 mm gap and a voltage of 1.8 kV was applied. The suspension was then mixed with 1 ml of R2A medium and incubated for 2 hr at 24°C with shaking. The transformants were selected on an R2A agar plate containing kanamycin (25 µg/ml).

Light and fluorescence microscopy

Cells were cultured in R2A medium either in a test tube containing liquid medium, or on a 1.5% agar or 3.0% gellan gum plate. The liquid-cultured cells were then observed on a glass slide under a microscope (Nikon Eclipse 80i, Nikon Inc, Japan). For time-lapse video microscopy, colonies grown on agar were observed directly on a Petri dish under the microscope using a ×10 or ×40 objective. Nile-red fluorescent images were obtained for the colonies that were inoculated and grown as described above. The colonies were observed under the microscope with filters for excitation (525–540 nm) and emission (>565 nm) wavelengths. To obtain three-dimensional images of a colony, a small piece of agar was cut out from an agar plate and placed on a glass slide. The slide was then observed under a confocal laser scanning microscope (Leica Microsystems TCS SP8, Leica Microsystems, Wetzlar, Germany) with filters for excitation (488–552 nm) and emission (560–620 nm) wavelengths and a ×10 objective. The images were processed using ImageJ 1.52 software (https://imagej.nih.gov/ij/).

Electron microscopy

A gel slice of a 48-hr-old colony was cut out from a Petri dish and soaked overnight in a 4% paraformaldehyde solution in 100 mM phosphate buffer (pH 7.4), and then soaked for 1 hr twice in 20 mM phosphate buffer (pH 7.0). The slice was then dried in a desiccator at room temperature overnight, and then dried under reduced pressure (−20 kPa) overnight. The slice was mounted on an appropriate grid and examined under a scanning electron microscope (JSM-6510LA, JEOL, Tokyo, Japan). Transmission electron microscopy (TEM) was performed at Hanaichi Ultrastructure Research Institute, Nagoya, Japan. The liquid-cultured cells were mounted on a grid and washed with distilled water at room temperature. Excess liquid was removed and the specimen was negatively stained using 2% uranyl acetate before being observed by TEM (JEM1200EX, JEOL).

Statistical analysis

All experiments were conducted in triplicate. Values represent the mean ± standard error of the mean of three independent experiments.

Acknowledgements

We thank the following people for their support with this study: Dr. Masahiro Nakano for discussion on aspects related to physics. Ms. Aya Ohta for isolating the bacterium, Ms. Satoko Nakanomori and Mr. Sakae Fukase for phenotype characterization, Mr. Hiroyuki Tanaka for SEM analysis, and Dr. Vishal Gor for his critical manuscript reading and English editing.

Funding Statement

No external funding was received for this work.

Contributor Information

Kouhei Mizuno, Email: [email protected].

Kazuya Morikawa, Email: [email protected].

Raymond E Goldstein, University of Cambridge, United Kingdom.

Detlef Weigel, Max Planck Institute for Biology Tübingen, Germany.

Additional information

Competing interests

No competing interests declared.

No competing interests declared.

Author contributions

Conceptualization, Data curation, Formal analysis, Supervision, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing.

Data curation, Formal analysis, Methodology, Writing – original draft, Writing – review and editing.

Investigation, Methodology, Writing – original draft, Writing – review and editing.

Investigation, Methodology.

Investigation, Methodology, Writing – original draft.

Conceptualization, Investigation, Methodology.

Investigation, Methodology, Writing – original draft.

Conceptualization, Supervision, Investigation, Methodology, Writing – original draft, Writing – review and editing.

Additional files

Supplementary file 1. Characteristics that distinguish strain HS-3 from its close phylogenetic relatives.
elife-71920-supp1.docx (25.4KB, docx)
Transparent reporting form

Data availability

The complete genome sequence and a plasmid sequence have been registered in the DDBJ database (https://www.ddbj.nig.ac.jp/) as AP024094 and AP024095, respectively.

The following previously published datasets were used:

Mizuno K, Maree M, Morikawa K. 2020. Jeongeupia sp. HS-3 plasmid pJHS3 DNA, complete sequence. NCBI Nucleotide. AP024095.1

Mizuno K, Maree M, Morikawa K. 2020. Jeongeupia sp. HS-3 DNA, complete genome. NCBI Nucleotide. AP024094.1

References

  1. Altschul SF, Madden TL, Schäffer AA, Zhang J, Zhang Z, Miller W, Lipman DJ. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Research. 1997;25:3389–3402. doi: 10.1093/nar/25.17.3389. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Amard B, Bertrand-Sarfati J. Microfossils in 2000 ma old cherty stromatolites of the franceville group, gabon. Precambrian Research. 1997;81:197–221. doi: 10.1016/S0301-9268(96)00035-6. [DOI] [Google Scholar]
  3. Barton HA, Northup DE. Geomicrobiology in cave environments: past, current and future perspectives. Journal of Cave and Karst Studies. 2007;69:163–178. [Google Scholar]
  4. Basu S, Gerchman Y, Collins CH, Arnold FH, Weiss R. A synthetic multicellular system for programmed pattern formation. Nature. 2005;434:1130–1134. doi: 10.1038/nature03461. [DOI] [PubMed] [Google Scholar]
  5. Bernardes JP, John U, Woltermann N, Valiadi M, Hermann RJ, Becks L. The evolution of convex trade-offs enables the transition towards multicellularity. Nature Communications. 2021;12:4222. doi: 10.1038/s41467-021-24503-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Black AJ, Bourrat P, Rainey PB. Ecological scaffolding and the evolution of individuality. Nature Ecology & Evolution. 2020;4:426–436. doi: 10.1038/s41559-019-1086-9. [DOI] [PubMed] [Google Scholar]
  7. Bonner JT. The origins of multicellularity. Integrative Biology. 1998;1:27–36. doi: 10.1002/(SICI)1520-6602(1998)1:1<27::AID-INBI4>3.0.CO;2-6. [DOI] [Google Scholar]
  8. Boraas ME, Seale DB, Boxhorn JE. Phagotrophy by A flagellate selects for colonial prey: A possible origin of multicellularity. Evolutionary Ecology. 1998;12:153–164. doi: 10.1023/A:1006527528063. [DOI] [Google Scholar]
  9. Brunet T, King N. The origin of animal multicellularity and cell differentiation. Developmental Cell. 2017;43:124–140. doi: 10.1016/j.devcel.2017.09.016. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Casadesús J, Low DA. Programmed heterogeneity: epigenetic mechanisms in bacteria. The Journal of Biological Chemistry. 2013;288:13929–13935. doi: 10.1074/jbc.R113.472274. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Cavalazzi B, Lemelle L, Simionovici A, Cady SL, Russell MJ, Bailo E, Canteri R, Enrico E, Manceau A, Maris A, Salomé M, Thomassot E, Bouden N, Tucoulou R, Hofmann A. Cellular remains in a ~3.42-billion-year-old subseafloor hydrothermal environment. Science Advances. 2021;7:eabf3963. doi: 10.1126/sciadv.abf3963. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Chen WM, Chang RC, Cheng CY, Shiau YW, Sheu SY. Jeongeupia chitinilytica sp. nov., a chitinolytic bacterium isolated from soil. International Journal of Systematic and Evolutionary Microbiology. 2013;63:934–938. doi: 10.1099/ijs.0.043125-0. [DOI] [PubMed] [Google Scholar]
  13. Claessen D, Rozen DE, Kuipers OP, Søgaard-Andersen L, van Wezel GP. Bacterial solutions to multicellularity: a tale of biofilms, filaments and fruiting bodies. Nature Reviews. Microbiology. 2014;12:115–124. doi: 10.1038/nrmicro3178. [DOI] [PubMed] [Google Scholar]
  14. Cowan ST, Steel KJ. Manual for the Identification of Medical Bacteria. London: Cambridge University Press; 1965. [Google Scholar]
  15. Dell’Arciprete D, Blow ML, Brown AT, Farrell FDC, Lintuvuori JS, McVey AF, Marenduzzo D, Poon WCK. A growing bacterial colony in two dimensions as an active nematic. Nature Communications. 2018;9:4190. doi: 10.1038/s41467-018-06370-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Doostmohammadi A, Ignés-Mullol J, Yeomans JM, Sagués F. Active nematics. Nature Communications. 2018;9:3246. doi: 10.1038/s41467-018-05666-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Dubnau D, Losick R. Bistability in bacteria. Molecular Microbiology. 2006;61:564–572. doi: 10.1111/j.1365-2958.2006.05249.x. [DOI] [PubMed] [Google Scholar]
  18. Duclos G, Erlenkämper C, Joanny JF, Silberzan P. Topological defects in confined populations of spindle-shaped cells. Nature Physics. 2017;13:58–62. doi: 10.1038/nphys3876. [DOI] [Google Scholar]
  19. Fukuyama M, Urata K, Nishiyama T. Geology and petrology of the hirao limestone and the tagawa metamorphic rocks-with special reference to the contact metamorphism by cretaceous granodiorite. Journal of Mineralogical and Petrological Sciences. 2004;99:25–41. doi: 10.2465/jmps.99.25. [DOI] [Google Scholar]
  20. Golubic S, Seong-Joo L. Early cyanobacterial fossil record: preservation, palaeoenvironments and identification. European Journal of Phycology. 2004;34:339–348. doi: 10.1080/09670269910001736402. [DOI] [Google Scholar]
  21. Hammerschmidt K, Rose CJ, Kerr B, Rainey PB. Life cycles, fitness decoupling and the evolution of multicellularity. Nature. 2014;515:75–79. doi: 10.1038/nature13884. [DOI] [PubMed] [Google Scholar]
  22. Herron MD, Borin JM, Boswell JC, Walker J, Chen ICK, Knox CA, Boyd M, Rosenzweig F, Ratcliff WC. De novo origins of multicellularity in response to predation. Scientific Reports. 2019;9:2328. doi: 10.1038/s41598-019-39558-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Ilham M, Nakanomori S, Kihara T, Hokamura A, Matsusaki H, Tsuge T, Mizuno K. Characterization of polyhydroxyalkanoate synthases from halomonas sp. O-1 and halomonas elongata DSM2581: site-directed mutagenesis and recombinant expression. Polymer Degradation and Stability. 2014;109:416–423. doi: 10.1016/j.polymdegradstab.2014.04.024. [DOI] [Google Scholar]
  24. Itcus C, Pascu MD, Lavin P, Perşoiu A, Iancu L, Purcarea C. Bacterial and archaeal community structures in perennial cave ice. Scientific Reports. 2018;8:15671. doi: 10.1038/s41598-018-34106-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Katayama-fujimura Y, Komatsu Y, Kuraishi H, Kaneko T. Estimation of DNA base composition by high performance liquid chromatography of its nuclease P1 hydrolysate. Agricultural and Biological Chemistry. 2014;48:3169–3172. doi: 10.1271/bbb1961.48.3169. [DOI] [Google Scholar]
  26. King N, Westbrook MJ, Young SL, Kuo A, Abedin M, Chapman J, Fairclough S, Hellsten U, Isogai Y, Letunic I, Marr M, Pincus D, Putnam N, Rokas A, Wright KJ, Zuzow R, Dirks W, Good M, Goodstein D, Lemons D, Li W, Lyons JB, Morris A, Nichols S, Richter DJ, Salamov A, Sequencing JGI, Bork P, Lim WA, Manning G, Miller WT, McGinnis W, Shapiro H, Tjian R, Grigoriev IV, Rokhsar D. The genome of the choanoflagellate monosiga brevicollis and the origin of metazoans. Nature. 2008;451:783–788. doi: 10.1038/nature06617. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Komagata K, Suzuki KI. Lipid and cell-wall analysis in bacterial Systematics. Methods in Microbiology. 1988;19:161–207. doi: 10.1016/S0580-9517(08)70410-0. [DOI] [Google Scholar]
  28. Koschwanez JH, Foster KR, Murray AW, Keller L. Sucrose utilization in budding yeast as a model for the origin of undifferentiated multicellularity. PLOS Biology. 2011;9:e1001122. doi: 10.1371/journal.pbio.1001122. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Kovach ME, Phillips RW, Elzer PH, Roop RM, Peterson KM. PBBR1MCS: a broad-host-range cloning vector. BioTechniques. 1994;16:800–802. [PubMed] [Google Scholar]
  30. Laiz L, Groth I, Gonzalez I, Saiz-Jimenez C. Microbiological study of the dripping waters in altamira cave (santillana del mar, spain) Journal of Microbiological Methods. 1999;36:129–138. doi: 10.1016/s0167-7012(99)00018-4. [DOI] [PubMed] [Google Scholar]
  31. Lane DJ. In: Nucleic Acid Techniques in Bacterial Systematic. Stackebrandt E, Goodfellow M, editors. New York: John Wiley and Sons; 1996. 16S/23S rrna sequencing; pp. 115–175. [Google Scholar]
  32. Lavoie KH, Winter AS, Read KJH, Hughes EM, Spilde MN, Northup DE. Comparison of bacterial communities from lava cave microbial mats to overlying surface soils from lava beds national monument, USA. PLOS ONE. 2017;12:e0169339. doi: 10.1371/journal.pone.0169339. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Libby E, Ratcliff WC. Evolution. ratcheting the evolution of multicellularity. Science. 2014;346:426–427. doi: 10.1126/science.1262053. [DOI] [PubMed] [Google Scholar]
  34. Lyons NA, Kolter R. On the evolution of bacterial multicellularity. Current Opinion in Microbiology. 2015;24:21–28. doi: 10.1016/j.mib.2014.12.007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Macalady JL, Dattagupta S, Schaperdoth I, Jones DS, Druschel GK, Eastman D. Niche differentiation among sulfur-oxidizing bacterial populations in cave waters. The ISME Journal. 2008;2:590–601. doi: 10.1038/ismej.2008.25. [DOI] [PubMed] [Google Scholar]
  36. Majumdar A, Cristina MM, Virga EG. Perspectives in active liquid crystals. Philosophical Transactions. Series A, Mathematical, Physical, and Engineering Sciences. 2014;372:20130373. doi: 10.1098/rsta.2013.0373. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Malik HS, Rosenberg SM. Life, death, differentiation, and the multicellularity of bacteria. PLOoS Genetics. 2009;5:e1000418. doi: 10.1371/journal.pgen.1000418. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Matt G, Umen J. Volvox: A simple algal model for embryogenesis, morphogenesis and cellular differentiation. Developmental Biology. 2016;419:99–113. doi: 10.1016/j.ydbio.2016.07.014. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Mizuno K, Ohta A, Hyakutake M, Ichinomiya Y, Tsuge T. Isolation of polyhydroxyalkanoate-producing bacteria from a polluted soil and characterization of the isolated strain bacillus cereus YB-4. Polymer Degradation and Stability. 2010;95:1335–1339. doi: 10.1016/j.polymdegradstab.2010.01.033. [DOI] [Google Scholar]
  40. Mizuno K, Kihara T, Tsuge T, Lundgren BR, Sarwar Z, Pinto A, Nomura CT. Cloning and heterologous expression of a novel subgroup of class IV polyhydroxyalkanoate synthase genes from the genus bacillus. Bioscience, Biotechnology, and Biochemistry. 2017;81:194–196. doi: 10.1080/09168451.2016.1230006. [DOI] [PubMed] [Google Scholar]
  41. Nadell CD, Bassler BL. A fitness trade-off between local competition and dispersal in Vibrio cholerae biofilms. PNAS. 2011;108:14181–14185. doi: 10.1073/pnas.1111147108. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Ortiz M, Neilson JW, Nelson WM, Legatzki A, Byrne A, Yu Y, Wing RA, Soderlund CA, Pryor BM, Pierson LS, Maier RM. Profiling bacterial diversity and taxonomic composition on speleothem surfaces in kartchner caverns, AZ. Microbial Ecology. 2013;65:371–383. doi: 10.1007/s00248-012-0143-6. [DOI] [PubMed] [Google Scholar]
  43. Rangseekaew P, Pathom-Aree W. Cave actinobacteria as producers of bioactive metabolites. Frontiers in Microbiology. 2019;10:387. doi: 10.3389/fmicb.2019.00387. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Richter DJ, Fozouni P, Eisen MB, King N. Gene family innovation, conservation and loss on the animal stem lineage. eLife. 2018;7:e34226. doi: 10.7554/eLife.34226. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Molecular Biology and Evolution. 1987;4:406–425. doi: 10.1093/oxfordjournals.molbev.a040454. [DOI] [PubMed] [Google Scholar]
  46. Sambrook J, Russel DW. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press; 2001. [Google Scholar]
  47. Sanchez T, Chen DTN, DeCamp SJ, Heymann M, Dogic Z. Spontaneous motion in hierarchically assembled active matter. Nature. 2012;491:431–434. doi: 10.1038/nature11591. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Sasser M. Identification of bacteria by gas chromatography of cellular fatty acids. MIDI Technical Note #101 1990
  49. Sauro F, Cappelletti M, Ghezzi D, Columbu A, Hong PY, Zowawi HM, Carbone C, Piccini L, Vergara F, Zannoni D, De Waele J. Microbial diversity and biosignatures of amorphous silica deposits in orthoquartzite caves. Scientific Reports. 2018;8:17569. doi: 10.1038/s41598-018-35532-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Schirrmeister BE, Antonelli A, Bagheri HC. The origin of multicellularity in cyanobacteria. BMC Evolutionary Biology. 2011;11:45. doi: 10.1186/1471-2148-11-45. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Sebé-Pedrós A, Ballaré C, Parra-Acero H, Chiva C, Tena JJ, Sabidó E, Gómez-Skarmeta JL, Di Croce L, Ruiz-Trillo I. The dynamic regulatory genome of capsaspora and the origin of animal multicellularity. Cell. 2016;165:1224–1237. doi: 10.1016/j.cell.2016.03.034. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Shapiro JA. Thinking about bacterial populations as multicellular organisms. Annual Review of Microbiology. 1998;52:81–104. doi: 10.1146/annurev.micro.52.1.81. [DOI] [PubMed] [Google Scholar]
  53. Smith R, Tan CM, Srimani JK, Pai A, Riccione KA, Song H, You LC. Programmed allee effect in bacteria causes a tradeoff between population spread and survival. PNAS. 2014;111:1969–1974. doi: 10.1073/pnas.1315954111. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Stackebrandt E, Lang E, Cousin S, Päuker O, Brambilla E, Kroppenstedt R, Lünsdorf H. Deefgea rivuli gen. nov., sp. nov., a member of the class betaproteobacteria. International Journal of Systematic and Evolutionary Microbiology. 2007;57:639–645. doi: 10.1099/ijs.0.64771-0. [DOI] [PubMed] [Google Scholar]
  55. Strahl H, Bürmann F, Hamoen LW. The actin homologue mreb organizes the bacterial cell membrane. Nature Communications. 2014;5:3442. doi: 10.1038/ncomms4442. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Thompson JD, Higgins DG, Gibson TJ. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Research. 1994;22:4673–4680. doi: 10.1093/nar/22.22.4673. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Trigos AS, Pearson RB, Papenfuss AT, Goode DL. Altered interactions between unicellular and multicellular genes drive hallmarks of transformation in a diverse range of solid tumors. PNAS. 2017;114:6406–6411. doi: 10.1073/pnas.1617743114. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Vlamakis H, Chai Y, Beauregard P, Losick R, Kolter R. Sticking together: building a biofilm the Bacillus subtilis way. Nature Reviews. Microbiology. 2013;11:157–168. doi: 10.1038/nrmicro2960. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Yaman YI, Demir E, Vetter R, Kocabas A. Emergence of active nematics in chaining bacterial biofilms. Nature Communications. 2019;10:2285. doi: 10.1038/s41467-019-10311-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Yoon JH, Choi JH, Kang SJ, Choi NS, Lee JS, Song JJ. Jeongeupia naejangsanensis gen. nov., sp. nov., a cellulose-degrading bacterium isolated from forest soil from naejang mountain in korea. International Journal of Systematic and Evolutionary Microbiology. 2010;60:615–619. doi: 10.1099/ijs.0.012591-0. [DOI] [PubMed] [Google Scholar]
  61. Zhou K, Zhang WY, Pan HM, Li JH, Yue HD, Xiao T, Wu LF. Adaptation of spherical multicellular magnetotactic prokaryotes to the geochemically variable habitat of an intertidal zone. Environmental Microbiology. 2013;15:1595–1605. doi: 10.1111/1462-2920.12057. [DOI] [PubMed] [Google Scholar]

Editor's evaluation

Raymond E Goldstein 1

This article reports the discovery of an unusual form of bacterial multicellularity – an organism that can exist in dense, filamentous multicellular structures and clusters of coccobacillus daughter cells. Experiments that mimic the periodic immersion that the bacteria experience in their natural cave environment suggest that water immersion plays a role in these life-cycle dynamics. This work, while rather qualitative, will likely attract great interest from a diverse range of scientists working on multicellularity, the biophysics of cell packing, and geobiological problems.

Decision letter

Editor: Raymond E Goldstein1

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Novel multicellular prokaryote discovered next to an underground stream" for consideration by eLife. Your article has been reviewed by 2 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Gisela Storz as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

1) The reviewers are enthusiastic about the discovery but think the description of its significance is lacking.

They suggest reframing it, not as an example of a missing 'group phase' (which isn't in fact missing, and this paper is not a very compelling argument in any case since it is not on a lineage that has ultimately evolved complex multicellularity), but rather as a new form of bacterial multicellularity.

'Ecological scaffolding' is a hot topic in the field- the idea that ecology gives rise to the first multicellular life cycles, which then are co-opted by developmental processes (see Black, Bourrat and Rainey, 2020). We suggest to focus on (a) the novel biology of a new type of bacterial multicellularity, which is significant since we know that bacteria were forming multicellular colonies back 3.42 billion years ago (Calvazzi 2021), with the first honest-to-goodness multicellular lineages (i.e., cyanobacteria), and (b) how reliable ecological cycles (ecological scaffolding) may have driven the evolution of a new type of bacterial multicellularity in these caves.

(2) The authors should place their work more clearly in the context of recent work on the advantages of multicellular forms in other evolutionary contexts, where response to environmental pressures are thought to drive adoption of certain strategies.

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Novel multicellular prokaryote discovered next to an underground stream" for further consideration by eLife. Your revised article has been evaluated by Gisela Storz (Senior Editor) and a Reviewing Editor. We apologize for the long delay in furnishing this decision.

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined below:

Summary of reviewers' comments:

The reviewers appreciate the changes you have made in response to the first round of review but are of the opinion that the way in which your discovery is framed is not particularly well justified and may even detract from the main message of your paper.

Our main concern is that the paper is framed about gaps in our knowledge about how multicellularity evolves, but the paper does not actually do a good job addressing these gaps. We don't actually have any information about what its unicellular ancestors looked like or behaved, how selection acted on early multicellular stages, what transitional steps in its evolution were like, etc. The authors cannot address these questions, because they just have a single contemporary species. And these questions have been addressed in a large literature that the authors do not cite- classically involving the volvocine green algae, and experimental evolution work in yeast, Chlamydomonas, and Pseudomonas.

What we do have in this paper is a very nice example of bacterial multicellularity that is different from other examples of bacterial multicellularity. And, importantly, it appears that multicellularity, in this case, is adaptive because of the periodic flooding behavior of the cave, which allows for groups to form, generate, and then periodically disperse. We thought that the environment in which these microbes presumably evolved in might provide some neat insight into their evolution, but even this claim must be made lightly since we do not know how these organisms actually evolved. Specifically, we suggested that periodic flooding might act as an ecological scaffold, creating an early life cycle in which groups form and disperse based purely on the abiotic environment. This connects nicely to recent theoretical work on scaffolding, but it seems from the paper that the authors' do not understand what scaffolding is or how their system can be interpreted in this context.

We think that the introduction needs to be reworked, as in its present form it appears as a meandering list of some of what is known about the early evolution of multicellularity, but it does not review the topic well, build to a critical point or identify gaps in the knowledge that the paper does a good job of addressing. This revised version is, in our opinion, not as good as the first one, as more information has been added but it just makes the arguments about the evolution of multicellularity less coherent.

In the same vein, we are concerned about Figure 1: first, ecological scaffolding is not widely accepted as a necessary step in the transition to multicellularity. In fact, we do not even have a single good example of it happening. That's actually why we suggested it for this paper in the last review, and we think that this paper might be the best example yet of an idea that so far has just been shown to work in models. Though, of course, we don't know if scaffolding played a role in the evolution of HS3, it seems plausible given the environment. Recurrent dynamism (which is their term for what ecological scaffolding is- these are the same exact argument!), dichotomic cell response, and trade-offs are not all hypothetical conditions for the origin of multicellularity. Scaffolding is, but dichotomic cell response and trade-offs are not conceptual foundations for how multicellularity evolves. A dichotomic cell response will usually be a trait that can be adaptive in a multicellular life cycle, usually because it provides a benefit through division of labor/differentiation. This is really just saying 'cellular differentiation' in using the authors' own unique terminology. And trade-offs are a part of life that constrains how evolution works- a universal law of nature that applies to all of evolutionary biology. Critically, these are three different kinds of ideas, one a conceptual scheme for how multicellularity evolves, one a description of how cells behave, and one a basic feature of all biological evolution.

We do not understand why they are in the figure as 'hypothetical conditions for the emergence of multicellularity'.

We believe that this paper stands on its own two feet purely as an example of fascinating organismal biology, a new kind of multicellular bacteria found in caves! That is important in its own right.

We thus urge the authors to go with their strengths. Talk about the discovery of a fascinating new multicellular organism. Perhaps include a bit about what this teaches us about evolution in the discussion. But to use this as a foundation for the paper appears to us as a mistake, as they fail to clearly identify the gaps in the knowledge of how their work addresses them.

eLife. 2022 Oct 11;11:e71920. doi: 10.7554/eLife.71920.sa2

Author response


Essential revisions:

1) The reviewers are enthusiastic about the discovery but think the description of its significance is lacking.

They suggest reframing it, not as an example of a missing 'group phase' (which isn't in fact missing, and this paper is not a very compelling argument in any case since it is not on a lineage that has ultimately evolved complex multicellularity), but rather as a new form of bacterial multicellularity.

'Ecological scaffolding' is a hot topic in the field- the idea that ecology gives rise to the first multicellular life cycles, which then are co-opted by developmental processes (see Black, Bourrat and Rainey, 2020). We suggest to focus on (a) the novel biology of a new type of bacterial multicellularity, which is significant since we know that bacteria were forming multicellular colonies back 3.42 billion years ago (Calvazzi 2021), with the first honest-to-goodness multicellular lineages (i.e., cyanobacteria), and (b) how reliable ecological cycles (ecological scaffolding) may have driven the evolution of a new type of bacterial multicellularity in these caves.

(2) The authors should place their work more clearly in the context of recent work on the advantages of multicellular forms in other evolutionary contexts, where response to environmental pressures are thought to drive adoption of certain strategies.

We have revised the Introduction, Discussion, and Figure 1 to appropriately contextualize this study as a new form of bacterial multicellularity (described in detail below).

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Summary of reviewers' comments:

The reviewers appreciate the changes you have made in response to the first round of review but are of the opinion that the way in which your discovery is framed is not particularly well justified and may even detract from the main message of your paper.

Our main concern is that the paper is framed about gaps in our knowledge about how multicellularity evolves, but the paper does not actually do a good job addressing these gaps. We don't actually have any information about what its unicellular ancestors looked like or behaved, how selection acted on early multicellular stages, what transitional steps in its evolution were like, etc. The authors cannot address these questions, because they just have a single contemporary species. And these questions have been addressed in a large literature that the authors do not cite- classically involving the volvocine green algae, and experimental evolution work in yeast, Chlamydomonas, and Pseudomonas.

We introduced and cited previous works in Introduction and Discussion and amended the Figure 1.

What we do have in this paper is a very nice example of bacterial multicellularity that is different from other examples of bacterial multicellularity. And, importantly, it appears that multicellularity, in this case, is adaptive because of the periodic flooding behavior of the cave, which allows for groups to form, generate, and then periodically disperse. We thought that the environment in which these microbes presumably evolved in might provide some neat insight into their evolution, but even this claim must be made lightly since we do not know how these organisms actually evolved. Specifically, we suggested that periodic flooding might act as an ecological scaffold, creating an early life cycle in which groups form and disperse based purely on the abiotic environment. This connects nicely to recent theoretical work on scaffolding, but it seems from the paper that the authors' do not understand what scaffolding is or how their system can be interpreted in this context.

We carefully considered the concept of ecological scaffolding in the context of this study, and reorganized the whole context of this manuscript to focus on the bacterial multicellularity evolved in the cave removing extra information as suggested by the reviewers.

We think that the introduction needs to be reworked, as in its present form it appears as a meandering list of some of what is known about the early evolution of multicellularity, but it does not review the topic well, build to a critical point or identify gaps in the knowledge that the paper does a good job of addressing. This revised version is, in our opinion, not as good as the first one, as more information has been added but it just makes the arguments about the evolution of multicellularity less coherent.

In the same vein, we are concerned about Figure 1: first, ecological scaffolding is not widely accepted as a necessary step in the transition to multicellularity. In fact, we do not even have a single good example of it happening. That's actually why we suggested it for this paper in the last review, and we think that this paper might be the best example yet of an idea that so far has just been shown to work in models. Though, of course, we don't know if scaffolding played a role in the evolution of HS3, it seems plausible given the environment. Recurrent dynamism (which is their term for what ecological scaffolding is- these are the same exact argument!), dichotomic cell response, and trade-offs are not all hypothetical conditions for the origin of multicellularity. Scaffolding is, but dichotomic cell response and trade-offs are not conceptual foundations for how multicellularity evolves. A dichotomic cell response will usually be a trait that can be adaptive in a multicellular life cycle, usually because it provides a benefit through division of labor/differentiation. This is really just saying 'cellular differentiation' in using the authors' own unique terminology. And trade-offs are a part of life that constrains how evolution works- a universal law of nature that applies to all of evolutionary biology. Critically, these are three different kinds of ideas, one a conceptual scheme for how multicellularity evolves, one a description of how cells behave, and one a basic feature of all biological evolution.

We do not understand why they are in the figure as 'hypothetical conditions for the emergence of multicellularity'.

We amended Figure 1 and removed some extra concepts in Introduction and Discussion.

Associated Data

    This section collects any data citations, data availability statements, or supplementary materials included in this article.

    Data Citations

    1. Mizuno K, Maree M, Morikawa K. 2020. Jeongeupia sp. HS-3 plasmid pJHS3 DNA, complete sequence. NCBI Nucleotide. AP024095.1
    2. Mizuno K, Maree M, Morikawa K. 2020. Jeongeupia sp. HS-3 DNA, complete genome. NCBI Nucleotide. AP024094.1

    Supplementary Materials

    Supplementary file 1. Characteristics that distinguish strain HS-3 from its close phylogenetic relatives.
    elife-71920-supp1.docx (25.4KB, docx)
    Transparent reporting form

    Data Availability Statement

    The complete genome sequence and a plasmid sequence have been registered in the DDBJ database (https://www.ddbj.nig.ac.jp/) as AP024094 and AP024095, respectively.

    The following previously published datasets were used:

    Mizuno K, Maree M, Morikawa K. 2020. Jeongeupia sp. HS-3 plasmid pJHS3 DNA, complete sequence. NCBI Nucleotide. AP024095.1

    Mizuno K, Maree M, Morikawa K. 2020. Jeongeupia sp. HS-3 DNA, complete genome. NCBI Nucleotide. AP024094.1


    Articles from eLife are provided here courtesy of eLife Sciences Publications, Ltd

    RESOURCES