Phenotypic Switching in Biofilm-Forming Marine BacteriumPaenibacillus lautus NE3B01

Neelam Mangwani • Supriya Kumari •

Sudhir K. Shukla • T. S. Rao • Surajit Das

Received: 18 November 2013 / Accepted: 28 November 2013 / Published online: 23 January 2014

� Springer Science+Business Media New York 2014

Abstract Biofilm-forming marine bacterium Paenibacillus

lautus NE3B01 was isolated from a mangrove ecosystem,

Odisha, India. This isolate formed a swarming type of colony

pattern on the solid culture medium with 0.5–2 % agar. Phase

contrast microscopy study of a growing colony of P. lautus on

solid media and swarming pattern revealed the existence of

two phenotypically distinct cells (i.e. cocci and rods) across

the colonies. However, in actively growing planktonic cul-

ture, only rod-shaped cells were observed. Biofilm growth

studies (crystal violet assay) with the isolate showed signifi-

cant biofilm formation by 6 h, and the detachment phase was

observed after 18 h. Biofilm parameters (such as total bio-

mass, roughness coefficient, biofilm thickness, etc.) of 24-h-

old P. lautus biofilm were studied by confocal scanning laser

microscopy (CSLM). The CSLM study showed that P. lautus

formed a biofilm with an average thickness of 14.8 ± 2.6 lm,

a high roughness coefficient (0.379 ± 0.103) and surface to

bio-volume ratio (4.59 ± 1.12 lm2/lm3), indicating a highly

uneven topography of the biofilm. This also indicates that the

24-h-old biofilm is in dispersal phase. Scanning electron

microphotographs of P. lautus also supported the existence of

two distinct phenotypes of P. lautus. The current findings

suggest that P. lautus has two vegetative phenotypes and to

decongest the overcrowded biofilm the bacterium can switch

over to motile rods from nonmotile cocci and vice versa.

Introduction

Bacteria tend to colonize when they encounter a solid

surface in aquatic environments. Colonization is a collec-

tive behaviour of bacterial cells that result in a swarming

pattern of growth and biofilm formation [32]. Multiple

factors together with intercellular and intracellular signal-

ling, extracellular polymers, cell surface appendages,

nutrition and environmental sensing are imperative aspects

that determine the spatial arrangement of cells in bacterial

colonies and biofilms [3, 4, 14]. Biofilms are ubiquitous in

environmental systems particularly in freshwater and

marine ecosystems [5]. Biofilm is a multicellular sessile

community of bacteria attached to a substratum and shel-

tered in a milieu of self-synthesised extracellular polymeric

substances [19, 20]. Apart from bacterial biofilms, the other

social aggregation that has received a great deal of interest

amongst researchers is the swarming motility. Many bac-

terial species undergo rapid and coordinated translocation

of their population across solid and semi-solid surface and

exhibit some intricate growth pattern known as swarming.

The swarming pattern formation represents multi-cellular

structure that provides protection and cooperation amongst

cells [9]. Swarming motility is a diffusion mechanism that

generates macroscopic patterns, which has been widely

studied in Pseudomonas, Bacillus, Vibrio, Escherichia,

Salmonella and Proteus [18]. In the recent time, some

Paenibacillus species (Paenibacillus vortex and P. den-

dritiformis) have been studied for their complex swarming

behaviour [1, 14].

Electronic supplementary material The online version of thisarticle (doi:10.1007/s00284-014-0525-8) contains supplementarymaterial, which is available to authorized users.

N. Mangwani � S. Kumari � S. Das (&)

Laboratory of Environmental Microbiology and Ecology

(LEnME), Department of Life Science, National Institute of

Technology, Rourkela 769 008, Odisha, India

e-mail: surajit@nitrkl.ac.in; surajit@myself.com

S. K. Shukla � T. S. Rao

Biofouling and Biofilm Processes Section, Water & Steam

Chemistry Division, BARC Facilities,

Kalpakkam 603 102, Tamil Nadu, India

123

Curr Microbiol (2014) 68:648–656

DOI 10.1007/s00284-014-0525-8

The strategies for survival (e.g. endospores and canni-

balism) ensure bacterial population protection when

encountered with intense environmental conditions or

nutrient stress [8, 11]. In some cases, bacteria also adopt

phenotypic heterogeneity (or bistability) [7]. Bacteria

multiply by cell division that yields genetically and mor-

phologically identical siblings. Conversely, it is also well

established that within an isogenic population there may be

a regulatory diversification. As a result, bacterial cells can

display various phenotypes. The phenotypic variation can

benefit bacteria, as far as their growth and survival are

concerned, with changing environmental conditions [10]. A

recent study by Be’er et al. [1] reported similar phenom-

enon in P. dendrtifoirmis colonies.

In the present study, we have used a combinational

microscopic approach to describe a cellular heterogeneity

and phenotypic switching in P. lautus micro-colony as well

as in biofilm. The bacterium showed to switch from non-

motile cocci to motile rods when overcrowding occurs in

the colony. The motile rods can switch back to cocci on

finding a nutrient-rich and open surface. To the best of our

knowledge, this is the first report on biofilm population

heterogeneity wherein P. lautus cells are coccoid in shape

in the centre of biofilm and at the periphery the cells are

transformed into motile rods and migrate to avoid

overcrowding.

Materials and Methods

Isolation and Characterization

Paenibacillus lautus NE3B01 was isolated from marine

water sample collected from Bhitarkanika mangrove eco-

system, Odisha, India. The strain was identified by 16S

rRNA gene sequencing, and phylogenetic tree was con-

structed following Dash et al. [5]. The culture was main-

tained in Luria–Bertani (LB) broth, and all the experiments

were performed in LB with 0.1 % of NaCl. The strain has

been submitted to Microbial Type Culture Collection and

Gene Bank, India (MTCC 11807).

Biofilm Screening and Growth Studies

Screening for biofilm formation was carried out by glass

tube assay following O’Toole et al. [24] and Jain et al. [16].

The tubes with attached cell mass visible after crystal

violet staining were considered as biofilm positive. Biofilm

growth of the isolate was monitored for 48 h using

microtiter plate assay. A colony was inoculated in LB broth

and incubated at 37 �C overnight. The strain was grown in

LB broth (37 �C/160 rpm); at log phase, cell mass was

harvested and washed with phosphate-buffered saline

(PBS) twice. It was resuspended in LB broth, and OD600nm

was adjusted to 0.5. 10 ll of this inoculum was transferred

to microtiter plate well with 190 ll of LB medium. At

regular time interval, the plate was washed with water

twice, air dried and stained with 0.2 % crystal violet for

5 min. Again, it was washed twice with water, air dried and

destained with 95 % ethanol for 30 min and OD595nm was

measured using 96 well plate reader (Multiplate reader,

PerkinElmer).

Swarming Motility

Swarming motility was evaluated by seeding 10 ll of

stationary phase culture mounted at the centre of 8-cm

diameter motility plates with 0.5, 1, 1.5 and 2 % LB agar

and incubated at 37 �C for 48–72 h. Swarm plates were

typically allowed to dry at room temperature under laminar

hood for few hours before being used. The swarming pat-

tern was observed under a transwhite illumination cham-

ber. 1.5 % LB agar swarming plate, for general cell

morphology and colony swarming pattern were observed

under inverted fluorescence microscope equipped with 40X

phase contrast objective lens (Olympus, 1 9 71)

Microscopy of Biofilm

A single colony was inoculated in LB broth and incubated at

37 �C, and the overnight grown culture was diluted 1:100 in

LB broth. 3 ml of the diluted suspension was transferred to

six-well cell culture plates. A sterile glass slide

(2 cm 9 2 cm) was placed in each well in order to grow the

biofilm. The plate was incubated at 37 �C for 24 h. After 24 h,

the slides were removed, gently washed with PBS and then

stained with 0.02 % acridine orange for 5 min and washed

again with PBS. Confocal scanning laser microscopy (CSLM)

studies were performed to monitor the biofilm architecture. A

thin cover slip was placed over the biofilms and mounted

upside down over the objective lens of CSLM (TCS-SP2-

AOBS) equipped with DM IRE 2-inverted microscope (Leica

Microsystems, Hessen, Wetzlar, Germany). About 10 image

stacks were collected randomly from different points in order

to get reliable data. Biofilm parameters were assayed as

described in Mangwani et al. [20] by analysing the image

stacks using COMSTAT [13]. Scanning electron microscopy

(SEM) of the P. lautus biofilm was carried by growing the

biofilms grown on glass slides (1 9 1 cm). The biofilm-

grown slides were fixed with 2.5 % glutaraldehyde (in PBS,

pH 7.2) at 4 �C for 12 h, washed with PBS and post fixed with

1 % aqueous solution of tannic acid. Later, the specimens

were dehydrated with 30, 70 and 100 % alcohol and coated

with platinum. The samples were analysed using scanning

electron microscope (Jeol T-330 Scanning Electron Micro-

scope, Germany). The planktonic cells were also stained with

N. Mangwani et al.: Phenotypic Switching in Paenibacillus lautus 649

123

0.2 % crystal violet, destained with 95 % ethanol for light

microscopy studies and observed under oil immersion lens

(1009).

Results and Discussion

Isolation and Characterization

The partial 1335 bp 16S rRNA gene sequence of the isolate

NE3B01 was submitted to NCBI GenBank under accession

number JX273779. The 16S rRNA gene analysis indicated

that the strain is phylogenetically related to genus Paeniba-

cillus. The sequence alignment was carried out using

CLUSTAL W tool in MEGA version 5. A phylogenetic tree

was constructed by the Neighbour Joining method based on

16S rRNA partial gene sequence (Supplementary Fig. S1).

The selected isolate was designated as P. lautus NE3B01.

Biofilm Screening and Growth Studies

Screening for biofilm formation was done by visualization of

attached cell mass to the glass tube visible after crystal violet

staining (Supplementary Fig. S2a). Biofilm is a surface-

associated growth, composed of cells and EPS that gets

stained by crystal violet [12]. Biofilm could easily be visually

seen after destaining with 95 % ethanol. Biofilm growth

pattern of P. lautus NE3B01 was monitored over a period of

48 h using the microtiter plate assay by measuring the crystal

violet absorbance at 595 nm. P. lautus NE3B01 biofilm

growth at different time intervals was recorded, the maximum

biofilm growth was observed after 6 h of growth and it was

monitored till 48th h (Fig. 1). From the biofilm growth study

of P. lautus NE3B01, it was noted that the bacterium possess a

typical biofilm formation and detachment phases. P. lautus

NE3B01 establishes mature biofilm within 6 h of growth

followed by a detachment phase that was observed from

12–13 h. The detachment phase was again followed by sec-

ondary biofilm formation phase. The secondary biofilm was

established within 6 h and started the dispersal phase after

7th h (see Fig. 1). The dispersal process enables bacteria to

colonize and reinitiate the biofilm development process [27].

A number of factors like cell density, nutrition, surface for

growth and physical conditions (temperature, pH, salinity and

sheer forces) are responsible for sustaining a biofilm. In

addition, gene expression pattern, cell signalling process and

quorum sensing are also key regulators that participate in a

biofilm development [16, 19, 22, 25].

Swarming Motility Assay

Figure 2a–d illustrates the swarming pattern of P. lautus

NE3B01 on the surface of solid medium with different

concentrations of agar (i.e. 0.5, 1, 1.5 and 2 %). The

swarming growth pattern was observed after 48–72 h. P.

lautus showed a complex swarming pattern even at higher

percentage of agar (i.e. up to 2 %). The swarming pattern

by the strain was characterized by discriminate vortices

(Fig. 2b–d). The vortex grew and expanded outwards with

distinct colonies at the end, while at tail region and bran-

ches there was no growth of any colony. However, on

softer surface (0.5 % agar), vortex formation was not dis-

tinct, but diffused growth without any colonies was

observed (Fig. 2a). Similar complex swarming pattern and

vortex formation have been studied in P. dendritiformis

and P. vortex, and it was concluded as a social event reg-

ulated by a number of environmental factors and flagella

movement [1, 14, 26]. Under laboratory conditions,

swarming motility is also dependent on the properties of

the surface bacteria, including the agar concentration and

the degree to which it is lubricated by the bacterium.

Bacterial swarming is a quick and harmonized migration of

cells across solid surface and mostly powered by flagella.

However, experimental studies with P. dendritiformis,

Pseudomonas aeruginosa and Bacillus subtilis have shown

that swarming patterns may also be due to the production

of surfactants that disperse the cells and initiate swarming

motility on solid surface [6, 9].

Phase contrast microscopy of P. lautus NE3B01 colony

The colony structure of P. lautus on 1.5 % agar swarming

plate was observed directly in a phase contrast inverted

microscope at different microscopic fields (Fig. 3a–f).

Figure 3b shows the magnified view of the colony and its

centre that has coccoid-shaped cells (Fig. 3a, zone 1).

Figure 3c shows a magnified view of a colony margin

without any rod-shaped cells (Fig. 3a, zone 1). Figure 3d

shows the magnified view of a colony close to other colony

having both cocci- and rod-shaped cells at the periphery

(Fig. 3a, zone 2). Figure 3e, f show the bacterial cells

Fig. 1 P. lautus NE3B01 biofilm growth at different time interval

(growth was quantified in terms of absorbance at 595 nm)

650 N. Mangwani et al.: Phenotypic Switching in Paenibacillus lautus

123

between colonies. Only rod-shaped cell is visible in the

region where fewer cocci can be seen in Fig. 3f (Fig 3a,

zone 3, 4). From the phase contrast microscopic study, it

was observed that the fraction of rod-shaped population

increased with increasing distance from the colony,

whereas the proportion of cocci was higher within colonies.

These observations suggest the presence of two distinct

phenotypes i.e. motile rods and non-motile cocci when

grown on a solid medium. Figure 3g shows an isolated

colony of P. lautus NE3B01 from which some motile rods

get detached from the colony and form a new colony

nearby (Fig. 3a, zone 5). It can be speculated that P. lautus

possesses a phenotypic switching i.e. transition from cocci

to rod under the condition of overcrowding. The rod gets

switched to cocci again on finding a new settlement surface

nearby where nutrients are sufficient and no competition is

present in close proximity. This speculation corroborates

with the study of Be’er et al. [1] where an inducible and

reversible phenotypic switching phenomenon was descri-

bed as a mechanism of survival by P. dendritiformis due to

overcrowding and sibling colony competition. P. dendrit-

iformis switch between rods and cocci by toxin and sig-

nalling proteins known as Slf and Ris (sibling lethal factor

and rod-inducing signals, respectively), which induce the

cocci to rod shape in response to local environmental

conditions. It might be possible that both P. dendritiformis

and P. lautus have some common regulator network

conserved in Paenibacillus genus responsible for the two

distinct phenotypes. However, from the current study on P.

lautus, the mechanism of transformation appears to be

different from what is reported in P. dendritiformis as no

spores were observed in the present study in the culture of

P. lautus NE3B01.

SEM and CSLM analysis of P. lautus NE3B01 biofilm

The CSLM images (see Fig. 4a, b) show the biofilm growth

of P. lautus NE3B01 on the glass surface after 24 h of

growth at 37 �C. Figure 4a shows a 24-h-old biofilm con-

sisting of many rod-shaped bacteria spaced interstitially

along with the mature micro-colonies. In contrast, Fig. 4b

illustrates a mature 24-h-old biofilm of P. lautus NE3B01

with a maximum biofilm thickness of *21 lm. Figure 3

shows that P. lautus NE3B01 forms a dense biofilm in a

distinct manner, having branched growth at the terminals

with vortices. A number of rod-shape single cells are

clearly visible in Fig. 4a. Table 1 illustrates the COM-

STAT analysis of P. lautus NE3B01 biofilm, which had an

average thickness of 14.82 ± 2.61 lm and maximum

thickness of 21.02 ± 2.10. P. lautus NE3B01 biofilm

showed high roughness coefficient (0.379 ± 0.103) and

surface to biovolume ratio (4.59 ± 1.12 lm2/lm3), indi-

cating either highly uneven topography of biofilm or dis-

persal event [30, 31]. In general, bacterial cells are

Fig. 2 Swarming pattern of P.

lautus NE3B01 at different

concentrations of agar a 0.5 %

agar showing bacterial mat,

b 1 % agar, c 1.5 % agar, d 2 %

agar. At higher concentration of

agar, vortices can be seen.

Diameter of vortices decreases

with increase in agar

percentage, whereas swarming

pattern becomes more distinct at

high concentration of agar

N. Mangwani et al.: Phenotypic Switching in Paenibacillus lautus 651

123

compactly entrapped in a biofilm, resulting in loss of

motility [16]. However, CSLM studies with P. lautus

NE3B01 biofilm manifest that a number of individual cells

were present in nearby biofilm (see Fig. 4b), which might

have dispersed from the parent biofilm. This suggests that

there was a detachment of cells resulting from dissociation

of biofilm. Thus, it can be hypothesized that P. lautus

NE3B01 forms a dynamic biofilm community with higher

cell detachment rate as compared to biofilms formed by

other bacteria.

Paenibacillus lautus NE3B01 biofilm and cell mor-

phology were also studied by high resolution SEM at dif-

ferent magnifications. Figure 5a, b show that P. lautus

NE3B01 formed a dense biofilm showing two distinct

phenotypes of cells. Both rod and coccoid cells were

observed in the biofilm. However, the shape of both rod-

and cocci-shaped cells was quite different from their typ-

ical morphology as reported in P. dendritiformis [1]. Some

cocci-shaped cells were more or less like a ring or bicon-

cave disc. The cocci cells were more in number compactly

Fig. 3 a–g Colony and cell morphology of P. lautus NE3B01

observed under phase contrast microscope. a Swarming pattern over

1.5 % agar. b Magnified view of colony and at its centre (a, zone 1).

c Magnified view of a colony margin (a, zone 1). d Two adjacent

colonies having both cocci and rod at the periphery (a, zone 2).

e Bacterial cell between colonies and only rod-shaped cell are visible

(a, zone 3). f At the place fewer cocci can be seen (a, zone 4). g An

isolated colony of P. lautus NE3B01 from which some motile rods get

detached from the colony and forming a new colony nearby (a, zone

5)

652 N. Mangwani et al.: Phenotypic Switching in Paenibacillus lautus

123

arranged and attached to the glass surface, while the rod-

shaped cell were present at the periphery of the biofilm.

Some appendages could easily be seen on rod cells

(Fig. 5b); the rod-shaped cell may represent the motile cell

getting detached from the biofilm as was also observed

under phase contrast microscope (See supplementary file

video 1). From the SEM images, it is inferred that phe-

notypic switching occurred in biofilm under overcrowding

condition to facilitate the cells to escape in planktonic form

and allow it to settle at a new place. Phase contrast

microscopy and SEM examination of actively growing

planktonic culture showed abundance of rod-shaped cells

(Fig. 5c, d). These findings support the occurrence of

phenotypic variation and bistable switching between two

phenotypes in P. lautus biofilm. In general, being a clone

population bacterial culture is believed to be homogenous.

However, differential expression of genes is possible

amongst bacterial subpopulations because of some random

fluctuations in transcription and translation [2, 28]. SEM

and phase contrast microscopy results indicate that P.

lautus cells have surface structures that can help in folding

and unfolding of cells that give perception of cocci- and

rod-shaped populations, which are not a truly different cell

population. These structures also help in twitching motility

over hard agar surface (supplementary file, video 1).

Bacteria may exhibit a wide variety of morphologies in

its growth phase; these changes have to be made to

maintain the consistency of a bacterial cell as per its sur-

rounding milieu. However, the consistency of a bacterial

shape could be affected due to environmental stress. To

cope with changing environment, adaptation via pheno-

typic heterogeneity is an imperative biological function.

The pleomorphism phenomenon is well established in

bacteria such as Helicobacter pylori, Deinococcus radio-

durans, Mycobacterium tuberculosis, Bacillus bifidus and

Staphylococcus aureus [21, 23, 33]. For instance, in D.

radiodurans, nutritional stress can alter its phenotype and

radiotolerance [17, 29]. Phenotypic heterogeneity helps to

optimize interactions amongst cells and with the surfaces to

which they attach. Cells in such communities possess

alterations in phenotype with respect to growth and gene

expression [15, 22]. In the present study, the biofilm

formed by the P. lautus showed coccoid cells in the middle

of the micro-colony whereas at the periphery most of the P.

lautus cells were rod shaped and were motile. It is specu-

lated that disc-like cells resembling a doughnut are an

intermediate stage between cocci- and rod-shaped mor-

photypes and transformation from cocci to rod may be the

stage that helps the detachment of cells from the biofilm. A

schematic diagram Fig. 6 explains hypothetically the dif-

ferent stages of P. lautus transformation. The capability of

P. lautus NE3B01 to survive as rod and coccoid morpho-

types comes into sight because of static growth condition

and increased population density. The bacterium acquires

the rod shape to disperse itself from the competitive

overcrowded colony and re-establishes itself on to a virgin

surface. The current findings on coexistence of two sub-

populations of P. lautus provide a distinctive feature of

Fig. 4 a Confocal scanning

laser microscopic image of P.

lautus NE3B01 after acridine

orange staining. Arrow pointing

the dispersing planktonic rod

cells around the biofilm. bMagnified view of P. lautus

NE3B01 biofilm

Table 1 Biofilm parameters of 24-h-old P. lautus NE3B01 biofilm

measured by CSLM

S.

no.

Biofilm parameters (Mean ± SE)

1. Total biomass (lm3/lm2) 10.81 ± 2.67

2. Roughness coefficient

(dimensionless)

0.379 ± 0.103

3. Surface area of biomass (lm2) 132401.6 ± 11083.39

4. Surface to biovolume ratio (lm2/

lm3)

4.59 ± 1.12

5. Maximum thickness (lm) 21.02 ± 2.10

6. Average thickness (lm) 14.82 ± 2.61

N. Mangwani et al.: Phenotypic Switching in Paenibacillus lautus 653

123

Fig. 5 a Scanning electron

microscopic image of P. lautus

NE3B01 biofilm, showing mat-

like compact structure.

b Differential cell morphology

of P. lautus NE3B01. Arrow

and circle pointing rod and

cocci cells, respectively. c Rod-

shaped planktonic cells after

crystal violet staining observed

under light microscope.

d Typical doughnut-shaped

cells of P. lautus NE3B01

observed under scanning

electron microscope

Fig. 6 A schematic diagram illustrating the different stages in phenotypic switching of P. lautus NE3B01 during biofilm growth

654 N. Mangwani et al.: Phenotypic Switching in Paenibacillus lautus

123

differential phenotype and bacterial adaptation to conges-

tion or overcrowding in a biofilm.

Conclusion

The present study reveals pleomorphism in P. lautus

NE3B01 micro-colonies and biofilm when grown on a solid

surface. Cells of the biofilm community switched from

non-motile cocci to motile rods due to competition for

space and survival. The strain showed typical phases of

biofilm formation and dispersal as well as swarming

motility on agar surface. This suggests that phenotypic

variation is reversible and is interconnected with cocci and

rods. Coccoid cells preferred to live in association, whereas

their transformation into rods results in acquisition of fla-

gellar motility and dispersal. The findings are suggestive of

mechanism to deal with space and nutrient scarcity for

long-term survival. The ability to replicate and maintain

coccoid forms and promote transformation to motile rods

requires further molecular investigations. However, in a

holistic view, the findings can be useful in understanding

bacterial adaptation and social behaviour while existing as

a colony or a biofilm.

Acknowledgments The authors would like to acknowledge the

authorities of NIT, Rourkela and BARC Facilities, Kalpakkam for

providing facilities. N.M. gratefully acknowledges the receipt of

fellowship from Ministry of Human Resource Development, Gov-

ernment of India for doctoral research. S.D. thanks the Department of

Biotechnology, Government of India for research grants on biofilm-

based enhanced bioremediation.

Conflicts of interest Authors of the manuscript declare no conflict

of interest.

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