**4. Air-liquid (A-L) interface biofilms**

In contrast to the archetypal S-L interface biofilms, bacterial biofilms also form at the airliquid (A-L) interface of static liquids and are sometimes referred to as 'pellicles' [30]. Perhaps the earliest experimental observations of these were made for *Bacterium aceti* and *B. xylinum* in 1886 [1, 40]. Both bacteria were isolated from beer undergoing acetic fermentation in which alcohol is converted into acetic acid. *B. aceti*, an acetic acid bacterium whose modern name is unclear, was found to produce a greasy-looking biofilm which varied in thickness from an 'almost invisible film' to a paper-thick structure depending on the growth medium [40]. In contrast, the *B. xylinum* isolate, which would probably now be recognised as a *Gluconacetobacter* spp. produced a 'vinegar plant' described as a jelly-like transparent mass at the bottom of the liquid, but under favourable conditions it could also produce a robust gelatinous A-L interface biofilm up to 25 mm thick [1].

4 Cellulose – Medical, Pharmaceutical and Electronic Applications

competition, and environmental stress [20, 22, 32-35].

and terminating with dispersal of mature biofilm communities.

**4. Air-liquid (A-L) interface biofilms** 

water-filled spaces.

and move away to colonise new surfaces in more favourable environments (reviewed in [22]). Biofilms of mixed bacterial communities and of individual species that develop on solid surfaces exposed to a continuous flow of nutrients form a thick layer generally described as consisting of differentiated mushroom and pillar-like structures separated by

A defining feature of many biofilms is the exopolysaccharide polymer 'slime' that encapsulate the bacteria and provide the main structural component or matrix of the biofilm [20, 22, 24-25]. Although generally assumed to be primarily composed of polysaccharides, e.g. alginate, PEL (a glucose-rich polymer) and PSL (a repeating pentasaccharide containing d-mannose, d-glucose and l-rhamnose) produced by *P. aeruginosa* PA01, PIA (a 28 kDa soluble linear (1-6)-N-acetylglucosamine) and related PNAG polymer produced by *Staphylococcus aureus* MN8m and *S. epidermidis* 13-1, and PIA-like polymers produced by *Escherichia coli* K-12 MG1655, biofilm matrices can also contain proteins and nucleic acids having significant structural roles (reviewed in [30]). Exopolysaccharides are typically viewed as a shared resource that provides a benefit to the biofilm community by maintaining structure, facilitating signalling, and protecting residents from predation,

A second characteristic common to many S-L interface biofilms has been the involvement of quorum sensing in micro-colony development, exopolysaccharide expression, and dispersal. For example, the quorum signalling molecule, acyl-homoserine lactone (AHL), functions as a signal for the development of *P. aeruginosa* PA01 and *Pseudomonas fluorescens* B52 biofilms [36-37]. However, mathematical models based on O2 and nutrient transport (diffusion) limitation result in similar biofilm architecture (reviewed in [38]), suggesting biofilm development is equally sensitive to environmental conditions as it may be to geneticallydetermined regulation. Although quorum sensing is important in the development of some biofilms, the bacterial community will exploit all available mechanisms to adapt to local environmental conditions. In order to further understand the development and role of biofilms, the local environment should be considered in terms of ecological landscape theory in which the spatial configuration of the biofilm biomass is shaped by multiple physical and biological factors [39]. It is therefore likely that biofilm formation is the net result of many independent interactions, rather than the result of a unique pathway initiating attachment

In contrast to the archetypal S-L interface biofilms, bacterial biofilms also form at the airliquid (A-L) interface of static liquids and are sometimes referred to as 'pellicles' [30]. Perhaps the earliest experimental observations of these were made for *Bacterium aceti* and *B. xylinum* in 1886 [1, 40]. Both bacteria were isolated from beer undergoing acetic fermentation in which alcohol is converted into acetic acid. *B. aceti*, an acetic acid bacterium whose modern name is unclear, was found to produce a greasy-looking biofilm which varied in thickness from an 'almost invisible film' to a paper-thick structure Vinegar plants are generally a consortia of acetic acid bacteria and yeasts which produce a zoogleal mat or mixed-species biofilm, and were traditionally used to produce vinegar from beer, cider or wine. Acetic fermentation is initiated by a starter culture known as the 'mother' and obtained from a previous vinegar in a process known as back-slopping [41]. A similar starter often referred to as a 'tea fungus' is used today to produce Kombucha, a carbonated cider-like drink from a sugary solution containing black tea (see the description given in [42]). Acetic acid bacteria, including *Gluconacetobacter* spp., can be isolated from these and similar consortia where they are responsible for the cellulose matrix-based biofilm (see an early review of the acetic acid bacteria by [43]). These artificially-maintained *Gluconacetobacter* spp. are probably better adapted to growth in static liquid conditions than environmental isolates recovered from rotting fallen fruit [44] and under the right conditions, some can produce a gelatinous 'plug' up to 20 mm deep in 10-12 days [45]. In these, cellulose expression and probably growth, is restricted to a thin 50-100 μm deep zone at the top, where it is limited by O2 diffusing from above and nutrients diffusing through the mature biofilm from below [45]. The growing biofilm is maintained in position by the accumulation of small CO2 bubbles and by pressing against the walls of the container as it develops.

We expect that smaller-scale A-L interface biofilms might also occur in a wide range of natural environments, such as the partially-saturated fluid-filled pore networks of soils, in temporary puddles collecting on plants and other surfaces after rainfall, water-logged leaf tissues, or in small protected bodies of water such as ponds where the surface is not disturbed by wind or currents. In these environments, biofilm development would be restricted by a combination of nutrient availability, O2 diffusion, physical disturbance, as well as microbial competition and predation by protists and nematodes.

A-L interface biofilms are readily produced in experimental static liquid-media microcosms [5, 11, 15], and an example of the *P. fluorescens* SBW25 Wrinkly Spreader A-L interface biofilm is shown in Figure 1. In a survey of environmental pseudomonads using nutrientrich liquid King's B microcosms, we categorised A-L interface biofilms on the basis of phenotype and physical robustness into the physically cohesive (PC), floccular mass (FM), waxy aggregate (WA) and viscous mass (VM)-class biofilms [15, 46]. The characteristics of these biofilm-types are summarised in Table 2 (see also Figure 2). A-L interface biofilm formation appears to be an evolutionary deep-rooted ability amongst bacteria, presumably with significant ecological advantages. In experimental microcosms, increases in competitive fitness of biofilm-formers have been observed compared to non-biofilm– forming strains, whilst the cost to being a biofilm-forming mutant in an environment not suited to these structures is also measurable [5, 47-49].



Cellulose Expression in *Pseudomonas fluorescens* SBW25 and Other Environmental Pseudomonads 7

Many aspects of the ecological and mechanistic bases of evolution have been investigated by the experimental evolution of bacteria (reviewed in [50-52]). The adaptive radiation of the soil and plant-associated pseudomonad, *P. fluorescens* SBW25 [10, 12], has been investigated in some detail using experimental King's B microcosms (see Figure 1) following the first report by Rainey and Travisano [11]. These can be incubated with shaking to provide a homogenous environment, or statically without physical disturbance to provide a heterogeneous environment. The initial wild-type SBW25 colonists of static microcosms rapidly establish a gradient in which O2 drops to < 0.05% of normal levels below a depth of 200μm [53]. This gradient produces heterogeneity in the microcosm and defines three niches for colonisation and adaptation: the A-L interface, the liquid column, and the vial bottom. In contrast, microcosms subject to constant and vigorous mixing do not develop an O2 gradient or different niches. Wild-type SBW25 rapidly radiates to produce a range of phenotypically distinguishable mutants (morphs or morphotypes) to occupy the different niches [11]. This diversification is reproducible and occurs rapidly, typically within ~100 generations and 1-3 days. The main morphotypes recovered from evolving populations of wild-type SBW25 in static microcosms include the Wrinkly Spreaders (WS) which produce a wrinkled colony morphology and colonise the A-L interface through the formation of a biofilm (Figure 1); the Smooth (SM) morphs, including wild-type SBW25, which produce round, smooth colonies and colonise the liquid column, and the Fuzzy Spreaders (FS) which are characterised by

fuzzy-topped colonies and colonise the anoxic bottom of static microcosms [11].

**6. Cellulose expression in** *P. fluorescens* **SBW25** 

unstable [5, 48-49].

In an effort to understand the mechanistic basis of the adaptive leap of wild-type SBW25 from the liquid column-colonising SM-morph to the WS A-L interface niche specialist, the underlying molecular biology of the WS phenotype was investigated. This work, described in the following section, ultimately showed that the evolutionary innovation was the use of cellulose to produce a physically robust and resilient biofilm which allowed the colonisation of the A-L interface. Competitive fitness experiments have demonstrated that the WS has a significant fitness advantage over non-biofilm–forming strains in static microcosms [5, 11, 49]. Simplistically, WS cells were able to intercept O2 diffusing across the A-L interface from the atmosphere before non-biofilm–forming competitors could do so lower down the liquid column, and as a result, WS populations could grow more rapidly than non-WS populations [53]. In contrast however, the WS do not enjoy a fitness advantage in shaken microcosms where the O2 concentrations are uniform or on agar plates where the WS phenotype is

In order to understand the underlying mechanistic basis of the WS phenotype, a minitransposon screening approach using mini-Tn*5* was adopted to identify critical genes and pathways [5]. Mini-transposon insertions typically destroy the function of the targeted gene, and the disruption of critical genes in the WS would be expected to result in mutants that produced rounded, smooth (SM)-like colonies rather than the typical WS colony. Plates

**5. Experimental evolution and the Wrinkly Spreader** 

Biofilm attributes compiled from [15, 46, 73]. Strength, ability to withstand weight applied to the top of the biofilm; Resilience, response to applied physical disturbance such as gentle or vigorous mixing; Attachment, connection to the microcosm vial walls in the meniscus region; Matrix, evidence of EPS from behaviour of samples during microscopy; Cellulose, evidence from Calcofluor-staining and fluorescent microscopy.

**Table 2.** Different classes of air-liquid (A-L) interface biofilms produced by environmental pseudomonads

**Figure 2. Air-liquid (A-L) interface biofilms.** A-L interface biofilms produced by environmental pseudomonads can be categorised into four biofilm types according to visual phenotype, robustness and resistance to physical disturbance. These are the **(A)** Physically cohesive (PC), **(B)** Floccular mass (FM), **(C)** Waxy aggregate (WA), and **(D)** Viscous mass (VM) types. Shown are biofilms in static King's B microcosms *(left)*, after pouring into petri dishes (*middle*), and after vigorous mixing (*right*). Figure adapted from [15].

## **5. Experimental evolution and the Wrinkly Spreader**

6 Cellulose – Medical, Pharmaceutical and Electronic Applications

Floccular mass

*Occurrence* Rare Common Common Common

*Strength* Strong Medium Strong Weak

Good, disruption produces flocs that are hard to

destroy

*Attachment* High Medium High Poor

**Table 2.** Different classes of air-liquid (A-L) interface biofilms produced by environmental

**Figure 2. Air-liquid (A-L) interface biofilms.** A-L interface biofilms produced by environmental pseudomonads can be categorised into four biofilm types according to visual phenotype, robustness and resistance to physical disturbance. These are the **(A)** Physically cohesive (PC), **(B)** Floccular mass (FM), **(C)** Waxy aggregate (WA), and **(D)** Viscous mass (VM) types. Shown are biofilms in static King's B microcosms *(left)*, after pouring into petri dishes (*middle*), and after vigorous mixing (*right*). Figure

Biofilm attributes compiled from [15, 46, 73]. Strength, ability to withstand weight applied to the top of the biofilm; Resilience, response to applied physical disturbance such as gentle or vigorous mixing; Attachment, connection to the microcosm vial walls in the meniscus region; Matrix, evidence of EPS from behaviour of samples during microscopy;

Physically cohesive

and elastic structure

Very good, hard to break into smaller

fragments

Observed Observed Observed

Viscous mass

Large viscous

Very poor, disruption solubilises the structure

(VM)

mass

(PC)

Multiple flocs Single-piece flexible

(FM)

Waxy aggregate

produces smaller

EPS, possible cellto-cell interactions

Cellulose, evidence from Calcofluor-staining and fluorescent microscopy.

fragments

(WA)

*Structure* Single-piece rigid and brittle structure

*Resilience* Good, disruption

*Matrix* No evidence for

pseudomonads

adapted from [15].

Many aspects of the ecological and mechanistic bases of evolution have been investigated by the experimental evolution of bacteria (reviewed in [50-52]). The adaptive radiation of the soil and plant-associated pseudomonad, *P. fluorescens* SBW25 [10, 12], has been investigated in some detail using experimental King's B microcosms (see Figure 1) following the first report by Rainey and Travisano [11]. These can be incubated with shaking to provide a homogenous environment, or statically without physical disturbance to provide a heterogeneous environment. The initial wild-type SBW25 colonists of static microcosms rapidly establish a gradient in which O2 drops to < 0.05% of normal levels below a depth of 200μm [53]. This gradient produces heterogeneity in the microcosm and defines three niches for colonisation and adaptation: the A-L interface, the liquid column, and the vial bottom. In contrast, microcosms subject to constant and vigorous mixing do not develop an O2 gradient or different niches. Wild-type SBW25 rapidly radiates to produce a range of phenotypically distinguishable mutants (morphs or morphotypes) to occupy the different niches [11]. This diversification is reproducible and occurs rapidly, typically within ~100 generations and 1-3 days. The main morphotypes recovered from evolving populations of wild-type SBW25 in static microcosms include the Wrinkly Spreaders (WS) which produce a wrinkled colony morphology and colonise the A-L interface through the formation of a biofilm (Figure 1); the Smooth (SM) morphs, including wild-type SBW25, which produce round, smooth colonies and colonise the liquid column, and the Fuzzy Spreaders (FS) which are characterised by fuzzy-topped colonies and colonise the anoxic bottom of static microcosms [11].

In an effort to understand the mechanistic basis of the adaptive leap of wild-type SBW25 from the liquid column-colonising SM-morph to the WS A-L interface niche specialist, the underlying molecular biology of the WS phenotype was investigated. This work, described in the following section, ultimately showed that the evolutionary innovation was the use of cellulose to produce a physically robust and resilient biofilm which allowed the colonisation of the A-L interface. Competitive fitness experiments have demonstrated that the WS has a significant fitness advantage over non-biofilm–forming strains in static microcosms [5, 11, 49]. Simplistically, WS cells were able to intercept O2 diffusing across the A-L interface from the atmosphere before non-biofilm–forming competitors could do so lower down the liquid column, and as a result, WS populations could grow more rapidly than non-WS populations [53]. In contrast however, the WS do not enjoy a fitness advantage in shaken microcosms where the O2 concentrations are uniform or on agar plates where the WS phenotype is unstable [5, 48-49].

## **6. Cellulose expression in** *P. fluorescens* **SBW25**

In order to understand the underlying mechanistic basis of the WS phenotype, a minitransposon screening approach using mini-Tn*5* was adopted to identify critical genes and pathways [5]. Mini-transposon insertions typically destroy the function of the targeted gene, and the disruption of critical genes in the WS would be expected to result in mutants that produced rounded, smooth (SM)-like colonies rather than the typical WS colony. Plates

containing hundreds or thousands of WS colonies could be easily screened for a few SM-like colonies which could then be isolated for further examination.

Cellulose Expression in *Pseudomonas fluorescens* SBW25 and Other Environmental Pseudomonads 9

MinD–like ATPase involved in the appropriate spatial localisation of

responsible for the polymerisation of UDP-Glucose into cellulose. Predicted integral transmembrane protein, contains conserved D residue, QXXRW, HAKAGN and QTP motifs, a PilZ domain. Binds

(originally thought to bind c-*di*-GMP). Often fused with WssB.

red colony staining, fluorescent microscopy, enzymatic digestion and structural analysis of

WssA (BcsQ, YhjQ) Cellulose synthase-associated positioning subunit (CAY46577.1):

WssB (BcsA, YhjO) Cellulose synthase subunit (CAY46578.1): catalytically-active subunit

WssC (BcsB, YhjN) Cellulose synthase subunit (CAY46579.1): unknown function

WssD (Orf1, YhjM) Cellulose synthase subunit (CAY46580.1): Endo-1,4-D-glucanase (D-

WssE (BcsC/S, YhjI) Cellulose synthase subunit (CAY46581.1): unknown function.

WssF (BcsX) Cellulose synthase-associated acetylation subunit (CAY46582.1):

WssG Cellulose synthase-associated acetylation subunit (CAY46583.1):

WssH Cellulose synthase-associated acetylation subunit (CAY46584.1):

WssI Cellulose synthase-associated acetylation subunit (CAY46585.1):

WssJ Cellulose synthase-associated positioning subunit (CAY46586.1):

*G. xylinus* Bcs and *E. coli* Yhj synonyms are provided in parentheses. Function suggested from Wss experiments and

The diazo dye, Congo red (CR), had been used previously to stain bacterial colonies expressing cellulose [61], and we used this technique to show that WS, WS-18, WS-6 and WS-9, but not WS-1, WS-13, WS-22, WS-15 and WS-25 mutants, appeared to express cellulose on King's B plates [5, 59]. WS and WS-18 biofilm material were subsequently stained with the more specific fluorescent dye, Calcofluor, and examined by fluorescent microscopy (Figure 4). This showed that the biofilm was dominated by an extensive network of extracellular cellulose, with fibres ranging from 0.02 μm to over 100 μm thick. In

suggested function is to present acyl groups to WssGHI.

AlgF-like protein involved in the acetylation of cellulose. Includes a

AlgI-like protein involved in the acetylation of cellulose. Predicted

AlgJ-like protein involved in the acetylation of cellulose. Localised to

MinD-like ATPase like WssA but apparently functionally redundant.

Includes a putative signal peptide.

integral transmembrane protein.

the cellulose synthase complex.

purified matrix material [5, 59-60].

Protein (synonyms) Function (Accession No.)

c-*di*-GMP.

family cellulase).

putative signal peptide.

the periplasm.

Wss homologue investigations. AlgFIJ homologues are from *P. aeruginosa* FRD1 [58]. **Table 3.** Predicted functions of the *Pseudomonas fluorescens* SBW25 Wss proteins

This approach allowed the identification and sequencing of the SBW25 *wss* operon containing ten genes (*wssA-J*) required for the WS phenotype and is shown in Figure 3 (*wss* is an acronym for WS structural locus, responsible for the production of the main structural component required for the WS phenotype) [5]. Overall, the *wss* operon showed strong similarity to the cellulose biosynthetic clusters originally identified as the *acs* operon in *Gluconacetobacter hansenii* (formerly *Acetobacter xylinus*) ATCC 23769 [54] and subsequently annotated as the *yhj* operon in the whole-genome sequence of *Escherichia coli* K-12 [55]. Most *acs* (*Acetobacter* cellulose-synthesizing) homologues are now referred to as *bcs* (bacterial cellulose synthesizing) genes as we do here (*yhj* has no meaning). The degree of homology between the *wss*, *bcs* and *yhj* genes at the amino acid level strongly suggested that the SBW25 *wss* operon encoded a functional cellulose synthase, and the predicted functions of the Wss proteins are listed in Table 3.

**Figure 3. Structure of the cellulose biosynthesis operon.** The *Pseudomonas fluorescens* SBW25 cellulose synthase is encoded by the *wss* operon (*wssA-J*, black arrows). The core synthase is composed of WssBCDE subunits and the associated acetylation activity produced by WssFGHI. WssA and WssJ may be involved in the correct cellular localisation of the Wss complex, though WssJ is functionally redundant. The locations of key mini-Tn*5* transposon insertions are indicated (open triangles). WS-1, 13, 22, 15 & 25 mutants are unable to express cellulose. WS-18, 9 & 6 mutants express un-acetylated cellulose. Upstream of the *wss* operon is tRNAThr and downstream a hypothetical protein of unknown function (grey arrows). Scale bar: 1 kb. Figure adapted from [5].

However, the SBW25 *wss* operon showed two notable differences to the *G. xylinus bcs* and *E. coli yhj* operons. First, the *wss* operon contains two MinD-like homologues, WssA and WssJ, not previously recognised as having a role in cellulose synthesis. WssJ shows 51% identity at the amino acid level with WssA, but only short sections of similarity at the nucleotide level and does not appear to be a simple repeat of the *wssA* gene sequence. As MinD is involved in cell division and determining cell polarity [56], WssA and WssJ were proposed to ensure the correct spatial localization of the cellulose synthase complex at the cell poles [5]. Subsequently, the WssA-homologue, YhjQ (BcsQ), has been shown to be essential for this in *E. coli* K-12 [57]. Secondly, the *wss* operon includes three genes, *wssGHI*, that shares homology with the alginate acetylation proteins of *P. aeruginosa* FRD1, AlgFIJ [58].

In order to demonstrate that the SBW25 *wss* operon encoded a functional cellulose synthase, and to determine the role of the alginate acetylation-like *wssGHI* genes, cellulose expression in the WS and mini-Tn*5* mutants was examined by a variety of techniques, including Congo red colony staining, fluorescent microscopy, enzymatic digestion and structural analysis of purified matrix material [5, 59-60].

8 Cellulose – Medical, Pharmaceutical and Electronic Applications

the Wss proteins are listed in Table 3.

colonies which could then be isolated for further examination.

containing hundreds or thousands of WS colonies could be easily screened for a few SM-like

This approach allowed the identification and sequencing of the SBW25 *wss* operon containing ten genes (*wssA-J*) required for the WS phenotype and is shown in Figure 3 (*wss* is an acronym for WS structural locus, responsible for the production of the main structural component required for the WS phenotype) [5]. Overall, the *wss* operon showed strong similarity to the cellulose biosynthetic clusters originally identified as the *acs* operon in *Gluconacetobacter hansenii* (formerly *Acetobacter xylinus*) ATCC 23769 [54] and subsequently annotated as the *yhj* operon in the whole-genome sequence of *Escherichia coli* K-12 [55]. Most *acs* (*Acetobacter* cellulose-synthesizing) homologues are now referred to as *bcs* (bacterial cellulose synthesizing) genes as we do here (*yhj* has no meaning). The degree of homology between the *wss*, *bcs* and *yhj* genes at the amino acid level strongly suggested that the SBW25 *wss* operon encoded a functional cellulose synthase, and the predicted functions of

**Figure 3. Structure of the cellulose biosynthesis operon.** The *Pseudomonas fluorescens* SBW25 cellulose synthase is encoded by the *wss* operon (*wssA-J*, black arrows). The core synthase is composed of WssBCDE subunits and the associated acetylation activity produced by WssFGHI. WssA and WssJ may

redundant. The locations of key mini-Tn*5* transposon insertions are indicated (open triangles). WS-1, 13, 22, 15 & 25 mutants are unable to express cellulose. WS-18, 9 & 6 mutants express un-acetylated cellulose. Upstream of the *wss* operon is tRNAThr and downstream a hypothetical protein of unknown

However, the SBW25 *wss* operon showed two notable differences to the *G. xylinus bcs* and *E. coli yhj* operons. First, the *wss* operon contains two MinD-like homologues, WssA and WssJ, not previously recognised as having a role in cellulose synthesis. WssJ shows 51% identity at the amino acid level with WssA, but only short sections of similarity at the nucleotide level and does not appear to be a simple repeat of the *wssA* gene sequence. As MinD is involved in cell division and determining cell polarity [56], WssA and WssJ were proposed to ensure the correct spatial localization of the cellulose synthase complex at the cell poles [5]. Subsequently, the WssA-homologue, YhjQ (BcsQ), has been shown to be essential for this in *E. coli* K-12 [57]. Secondly, the *wss* operon includes three genes, *wssGHI*, that shares

be involved in the correct cellular localisation of the Wss complex, though WssJ is functionally

homology with the alginate acetylation proteins of *P. aeruginosa* FRD1, AlgFIJ [58].

In order to demonstrate that the SBW25 *wss* operon encoded a functional cellulose synthase, and to determine the role of the alginate acetylation-like *wssGHI* genes, cellulose expression in the WS and mini-Tn*5* mutants was examined by a variety of techniques, including Congo

function (grey arrows). Scale bar: 1 kb. Figure adapted from [5].


*G. xylinus* Bcs and *E. coli* Yhj synonyms are provided in parentheses. Function suggested from Wss experiments and Wss homologue investigations. AlgFIJ homologues are from *P. aeruginosa* FRD1 [58].

**Table 3.** Predicted functions of the *Pseudomonas fluorescens* SBW25 Wss proteins

The diazo dye, Congo red (CR), had been used previously to stain bacterial colonies expressing cellulose [61], and we used this technique to show that WS, WS-18, WS-6 and WS-9, but not WS-1, WS-13, WS-22, WS-15 and WS-25 mutants, appeared to express cellulose on King's B plates [5, 59]. WS and WS-18 biofilm material were subsequently stained with the more specific fluorescent dye, Calcofluor, and examined by fluorescent microscopy (Figure 4). This showed that the biofilm was dominated by an extensive network of extracellular cellulose, with fibres ranging from 0.02 μm to over 100 μm thick. In

places, the fibres appeared to form large clumps of material and in other places forming thin films, with bacterial cells associated with the fibres and found within the voids [59]. In comparison, in colonies the cellulose fibres appear to collect above the mass of the colony (Figure 5). Scanning electron microscopy images of WS biofilms suggest a lattice-work of pores (Figure 6) which might be the result of constant growth at the top surface of the biofilm which slowly displaces older strata deeper into the liquid column [53]. Rough calculations of the density of WS biofilms suggest that they were > 97% liquid, which is in agreement with the finding that microbial amorphous celluloses are very hydrophilic, with gels having a water holding capacity of 148 – 309 g water / g dry cellulose [62-63]. Recent rheological tests have shown that the WS biofilm structure is a classic viscoelastic solid (gellike) material (AK & AJS, unpublished observations). The structural integrity of WS biofilms could be destroyed by incubation with cellulase, adding support to initial conclusions that the major matrix component of the WS biofilm, expressed by the *wss* operon, was cellulose or a cellulose-like polymer [5, 59].

Cellulose Expression in *Pseudomonas fluorescens* SBW25 and Other Environmental Pseudomonads 11

identified a major peak corresponding to 4-Glu, and minor peaks corresponding to 2,4-Glu, 3,4-Glu and 4,6-Glu, which is consistent with a (1-4)-linked glucose polymer, i.e. cellulose. In contrast, the WS-18 material did not contain 2,4-Glu, 3,4-Glu or 4,6-Glu derivatives, suggesting that the *wss* alginate acetylation-like genes were responsible for the acetylation of glucose residues at the 2, 3, and 6 Carbon positions. This was further supported by [1H]- NMR analysis which confirmed the presence of acetylated hexose residues in the WS extract, with 14% of the glucose residues estimated to be modified with one acetyl group [59]. Although cellulose is readily acetylated by chemical treatment, we are not aware of any

**Figure 5. Inducing cellulose expression with c-***di***-GMP.** An increase in c-*di*-GMP levels can induce cellulose expression in some pseudomonads. Shown are confocal laser scanning microscopy (CLSM) images of colony material from **(A)** *Pseudomonas fluorescens* SBW25 and **(B)** *P. syringae* DC3000

expressing the constitutively-active DGC response regulator WspR19 *in trans* which increases c-*di*-GMP levels, visualised with Calcofluor for cellulose (blue) and ethidium bromide for bacteria (red). Scale bar:

The partial acetylation of the cellulose fibres expressed by the WS clearly had an impact on colony morphology and biofilm strength. Colonies produced by WS-18 were readily differentiated from WS and wild-type SM-like colonies, whilst the WS-18 biofilm was ~ 4*x* weaker than the partially-acetylated structure produced by the WS, suggesting that acetylation increased the connectivity of cellulose fibres within colonies and the biofilm matrix [59-60]. Incubation with EDTA reduced WS-18 biofilm strength, whilst incubation with some diazo dyes increased WS-18 biofilm strength compared to WS biofilms, suggesting that fibre interactions could be altered by sequestering Mg2+ and coating cellulose fibres with dyes [60]. A lipopolysaccharide-deficient mutant which produces a very weak biofilm compared to both WS-18 and WS, was also affected by EDTA and diazo dyes, indicating that the partially-acetylated cellulose fibres also interacted with the lipopolysaccharide on the surface of cells or associated with cell debris to further strengthen

other reports of biologically-produced acetylated cellulose.

10 μm. Images from O. Moshynets.

the biofilm structure [60].

**Figure 4. Fluorescent microscopy of WS biofilms.** The cellulose fibre matrix of the Wrinkly Spreader (WS) biofilm can be visualised by staining with Calcofluor and fluorescent microscopy. Shown are two images showing the highly hydrated and fibrous nature of the WS biofilm. Scale bar: 100 μm. Images from A. Spiers.

WS and WS-18 biofilms were subsequently purified in order to determine the chemical identity of the matrix component. Carbohydrate analysis indicated that both samples contained ~75% glucose (Glu) and ~25% rhamnose (Rha) [59]. The latter could be explained as coming from contaminating Rha-containing A-band LPS which is highly conserved amongst the pseudomonads [64]. Linkage analyses of derivatized WS samples by GC-MS identified a major peak corresponding to 4-Glu, and minor peaks corresponding to 2,4-Glu, 3,4-Glu and 4,6-Glu, which is consistent with a (1-4)-linked glucose polymer, i.e. cellulose. In contrast, the WS-18 material did not contain 2,4-Glu, 3,4-Glu or 4,6-Glu derivatives, suggesting that the *wss* alginate acetylation-like genes were responsible for the acetylation of glucose residues at the 2, 3, and 6 Carbon positions. This was further supported by [1H]- NMR analysis which confirmed the presence of acetylated hexose residues in the WS extract, with 14% of the glucose residues estimated to be modified with one acetyl group [59]. Although cellulose is readily acetylated by chemical treatment, we are not aware of any other reports of biologically-produced acetylated cellulose.

10 Cellulose – Medical, Pharmaceutical and Electronic Applications

or a cellulose-like polymer [5, 59].

from A. Spiers.

places, the fibres appeared to form large clumps of material and in other places forming thin films, with bacterial cells associated with the fibres and found within the voids [59]. In comparison, in colonies the cellulose fibres appear to collect above the mass of the colony (Figure 5). Scanning electron microscopy images of WS biofilms suggest a lattice-work of pores (Figure 6) which might be the result of constant growth at the top surface of the biofilm which slowly displaces older strata deeper into the liquid column [53]. Rough calculations of the density of WS biofilms suggest that they were > 97% liquid, which is in agreement with the finding that microbial amorphous celluloses are very hydrophilic, with gels having a water holding capacity of 148 – 309 g water / g dry cellulose [62-63]. Recent rheological tests have shown that the WS biofilm structure is a classic viscoelastic solid (gellike) material (AK & AJS, unpublished observations). The structural integrity of WS biofilms could be destroyed by incubation with cellulase, adding support to initial conclusions that the major matrix component of the WS biofilm, expressed by the *wss* operon, was cellulose

**Figure 4. Fluorescent microscopy of WS biofilms.** The cellulose fibre matrix of the Wrinkly Spreader (WS) biofilm can be visualised by staining with Calcofluor and fluorescent microscopy. Shown are two images showing the highly hydrated and fibrous nature of the WS biofilm. Scale bar: 100 μm. Images

WS and WS-18 biofilms were subsequently purified in order to determine the chemical identity of the matrix component. Carbohydrate analysis indicated that both samples contained ~75% glucose (Glu) and ~25% rhamnose (Rha) [59]. The latter could be explained as coming from contaminating Rha-containing A-band LPS which is highly conserved amongst the pseudomonads [64]. Linkage analyses of derivatized WS samples by GC-MS

**Figure 5. Inducing cellulose expression with c-***di***-GMP.** An increase in c-*di*-GMP levels can induce cellulose expression in some pseudomonads. Shown are confocal laser scanning microscopy (CLSM) images of colony material from **(A)** *Pseudomonas fluorescens* SBW25 and **(B)** *P. syringae* DC3000 expressing the constitutively-active DGC response regulator WspR19 *in trans* which increases c-*di*-GMP levels, visualised with Calcofluor for cellulose (blue) and ethidium bromide for bacteria (red). Scale bar: 10 μm. Images from O. Moshynets.

The partial acetylation of the cellulose fibres expressed by the WS clearly had an impact on colony morphology and biofilm strength. Colonies produced by WS-18 were readily differentiated from WS and wild-type SM-like colonies, whilst the WS-18 biofilm was ~ 4*x* weaker than the partially-acetylated structure produced by the WS, suggesting that acetylation increased the connectivity of cellulose fibres within colonies and the biofilm matrix [59-60]. Incubation with EDTA reduced WS-18 biofilm strength, whilst incubation with some diazo dyes increased WS-18 biofilm strength compared to WS biofilms, suggesting that fibre interactions could be altered by sequestering Mg2+ and coating cellulose fibres with dyes [60]. A lipopolysaccharide-deficient mutant which produces a very weak biofilm compared to both WS-18 and WS, was also affected by EDTA and diazo dyes, indicating that the partially-acetylated cellulose fibres also interacted with the lipopolysaccharide on the surface of cells or associated with cell debris to further strengthen the biofilm structure [60].

Individual cellulose polymers can also interact directly to produce a number of different forms or allomorphs. *G. xylinus* produces two crystalline allomorphs, known as cellulose I and II, which requires the cellulose synthase-associated BcsD subunit that couples cellulose polymerisation and crystallization [54, 65]. However, SBW25 lacks a BcsD homologue and therefore can only produce non-crystalline amorphous cellulose.

Cellulose Expression in *Pseudomonas fluorescens* SBW25 and Other Environmental Pseudomonads 13

Examination of other mini-Tn*5* mutants also lead to the identification of the *wsp* regulatory operon of seven genes, *wspA-E & R* (*wsp* is an acronym for WS phenotype locus, responsible for the regulation of the WS phenotype) [67]. The function of these have been modelled on the *Escherichia coli* Che chemosensory system (reviewed in [68]) to provide a mechanistic explanation of the induction of the WS phenotype [67] (a schematic of this is shown in Figure 7). In this the methyl-accepting chemotaxis protein (MCP) WspA forms a membranebound complex with two scaffold proteins, WspB and WspD, plus the histidine kinase WspE. In the absence of an appropriate environmental signal, the complex is silent and does not activate the associate response regulator, WspR, by phosphorylation. The system is controlled by a negative feedback loop mediated by the WspC methyltransferase and WspF methylesterase. WspC constitutively antagonises WspF, and in wild-type SBW25 the activities of the two are balanced, preventing the activation of WspR and allowing the Wsp complex to oscillate between active and inactive states. WspR is a di-guanylate cyclase (DGC) response regulator, and the phosphorylated active form, WspR-P, synthesizes c-*di*-GMP (bis-(3'-5')-cyclic dimeric guanosine monophosphate) from GTP [69]. In this model, we hypothesised that mutations in WspR which stimulated DGC activity without requiring phosphorylation, or mutations inhibiting WspF function, would result in an increase in c-*di*-GMP production. This would then lead to the activation of the WS phenotype through the direct stimulation of the cellulose synthase complex, rather than up-regulated *wss* transcription [5] (the second component required for the WS phenotype, the pilli-like attachment factor, has not yet been identified and it is not known how it might be regulated by c-*di*-GMP). Several mutants of WspR had been engineered, and the effect of the constitutively-active mutant

WspR19 [70] on cellulose expression by wild-type SBW25 is shown in Figure 5.

leading to new A-L interface biofilm-forming genotypes in static microcosms [72].

synthase complex itself that allows activation despite sub-critical levels of c-*di*-GMP.

This model for the activation of the WS phenotype has been confirmed through the identification and testing of WspF mutations found in a number of independently-isolated Wrinkly Spreaders [67]. Interestingly, no naturally occurring WspR mutants have been identified yet, despite the fact that engineered WspR mutants like WspR19 were found to show the predicted phenotype [60, 69-71]. Mutations in other operons leading to the activation of the AwsR and MwsR DGCs can also induce the WS phenotype [47, 72]. These different routes activating the WS phenotype can be seen as an example of parallel evolution

During the molecular investigation of the WS phenotype, the non-biofilm–forming wildtype SBW25 was modified by the insertion of a constitutive promoter to increase the levels of *wss* operon transcription. This mutant, JB01, was found to produce a very weak biofilm, poorly attached to the microcosm vial walls and to express similar amounts of cellulose as the WS [5, 60]. Subsequently, we found that wild-type SBW25 could be non-specifically induced by exogenous Fe to produce a phenotypically-similar biofilm referred to as the VM biofilm [73] (see below for a description of Viscous mass (VM)-class biofilms). However, VM biofilm-formation was not the result of an increase in *wss* transcription, and as yet, no link has been identified between Fe regulation and cellulose expression. We speculate that the induction of the VM biofilm is due to a minor perturbation of the Wsp system or the cellulose

**Figure 6. WS biofilm ultrastructure.** Scanning electron microscopy (SEM) images of Wrinkly Spreader (WS) biofilms suggest a porous but robust structure. Shown here are a series of SEM images of decreasing magnification, from **(A)** single cells to **(F)** large pieces of biofilm. Images were obtained after freeze-drying and shadowing with gold. Scale bars: A & B, 1μm; C – F, 10 μm. Images from O. Moshynets.

Following the analysis of the WS mini-Tn*5* mutants, a further round of mini-transposon mutagenesis was undertaken using IS*phoA*/hah [66]. This mini-transposon allowed both the impact of polar and non-polar insertions to be assessed; the former destroy the function of the disrupted gene as well as any down-stream expression, whilst the latter destroys gene function but leaves down-stream expression unaffected (this is possible after Cre-mediated deletion of the central portion of the IS*phoA*/hah cassette which leaves a 63-codon in-frame insertion). WS IS*phoA*/hah mutants were recovered for each of the *wss* genes, except *wssJ*. Each of the polar IS*phoA*/hah insertions and corresponding non-polar Cre-deletions in *wssA-E* resulted in the loss of cellulose expression, whilst polar and non-polar mutants in *wssF-I*  resulted in a WS-18–like phenotype [47, 59]. Finally, a WS *wssJ* deletion mutant was constructed and shown to have no impact on cellulose expression, suggesting that the final gene of the *wss* operon might be functionally redundant [47].

therefore can only produce non-crystalline amorphous cellulose.

Individual cellulose polymers can also interact directly to produce a number of different forms or allomorphs. *G. xylinus* produces two crystalline allomorphs, known as cellulose I and II, which requires the cellulose synthase-associated BcsD subunit that couples cellulose polymerisation and crystallization [54, 65]. However, SBW25 lacks a BcsD homologue and

**Figure 6. WS biofilm ultrastructure.** Scanning electron microscopy (SEM) images of Wrinkly Spreader

decreasing magnification, from **(A)** single cells to **(F)** large pieces of biofilm. Images were obtained after

Following the analysis of the WS mini-Tn*5* mutants, a further round of mini-transposon mutagenesis was undertaken using IS*phoA*/hah [66]. This mini-transposon allowed both the impact of polar and non-polar insertions to be assessed; the former destroy the function of the disrupted gene as well as any down-stream expression, whilst the latter destroys gene function but leaves down-stream expression unaffected (this is possible after Cre-mediated deletion of the central portion of the IS*phoA*/hah cassette which leaves a 63-codon in-frame insertion). WS IS*phoA*/hah mutants were recovered for each of the *wss* genes, except *wssJ*. Each of the polar IS*phoA*/hah insertions and corresponding non-polar Cre-deletions in *wssA-E* resulted in the loss of cellulose expression, whilst polar and non-polar mutants in *wssF-I*  resulted in a WS-18–like phenotype [47, 59]. Finally, a WS *wssJ* deletion mutant was constructed and shown to have no impact on cellulose expression, suggesting that the final

(WS) biofilms suggest a porous but robust structure. Shown here are a series of SEM images of

freeze-drying and shadowing with gold. Scale bars: A & B, 1μm; C – F, 10 μm. Images from O.

gene of the *wss* operon might be functionally redundant [47].

Moshynets.

Examination of other mini-Tn*5* mutants also lead to the identification of the *wsp* regulatory operon of seven genes, *wspA-E & R* (*wsp* is an acronym for WS phenotype locus, responsible for the regulation of the WS phenotype) [67]. The function of these have been modelled on the *Escherichia coli* Che chemosensory system (reviewed in [68]) to provide a mechanistic explanation of the induction of the WS phenotype [67] (a schematic of this is shown in Figure 7). In this the methyl-accepting chemotaxis protein (MCP) WspA forms a membranebound complex with two scaffold proteins, WspB and WspD, plus the histidine kinase WspE. In the absence of an appropriate environmental signal, the complex is silent and does not activate the associate response regulator, WspR, by phosphorylation. The system is controlled by a negative feedback loop mediated by the WspC methyltransferase and WspF methylesterase. WspC constitutively antagonises WspF, and in wild-type SBW25 the activities of the two are balanced, preventing the activation of WspR and allowing the Wsp complex to oscillate between active and inactive states. WspR is a di-guanylate cyclase (DGC) response regulator, and the phosphorylated active form, WspR-P, synthesizes c-*di*-GMP (bis-(3'-5')-cyclic dimeric guanosine monophosphate) from GTP [69]. In this model, we hypothesised that mutations in WspR which stimulated DGC activity without requiring phosphorylation, or mutations inhibiting WspF function, would result in an increase in c-*di*-GMP production. This would then lead to the activation of the WS phenotype through the direct stimulation of the cellulose synthase complex, rather than up-regulated *wss* transcription [5] (the second component required for the WS phenotype, the pilli-like attachment factor, has not yet been identified and it is not known how it might be regulated by c-*di*-GMP). Several mutants of WspR had been engineered, and the effect of the constitutively-active mutant WspR19 [70] on cellulose expression by wild-type SBW25 is shown in Figure 5.

This model for the activation of the WS phenotype has been confirmed through the identification and testing of WspF mutations found in a number of independently-isolated Wrinkly Spreaders [67]. Interestingly, no naturally occurring WspR mutants have been identified yet, despite the fact that engineered WspR mutants like WspR19 were found to show the predicted phenotype [60, 69-71]. Mutations in other operons leading to the activation of the AwsR and MwsR DGCs can also induce the WS phenotype [47, 72]. These different routes activating the WS phenotype can be seen as an example of parallel evolution leading to new A-L interface biofilm-forming genotypes in static microcosms [72].

During the molecular investigation of the WS phenotype, the non-biofilm–forming wildtype SBW25 was modified by the insertion of a constitutive promoter to increase the levels of *wss* operon transcription. This mutant, JB01, was found to produce a very weak biofilm, poorly attached to the microcosm vial walls and to express similar amounts of cellulose as the WS [5, 60]. Subsequently, we found that wild-type SBW25 could be non-specifically induced by exogenous Fe to produce a phenotypically-similar biofilm referred to as the VM biofilm [73] (see below for a description of Viscous mass (VM)-class biofilms). However, VM biofilm-formation was not the result of an increase in *wss* transcription, and as yet, no link has been identified between Fe regulation and cellulose expression. We speculate that the induction of the VM biofilm is due to a minor perturbation of the Wsp system or the cellulose synthase complex itself that allows activation despite sub-critical levels of c-*di*-GMP.

Cellulose Expression in *Pseudomonas fluorescens* SBW25 and Other Environmental Pseudomonads 15

Calcofluor-fluorescent microscopy identified cellulose as the matrix component of 20% of the biofilm-forming isolates, indicating that at least seven *Pseudomonas* species were capable of expressing cellulose under the conditions tested. These included *P. corrugata* (tomato pathogens), *P. fluorescens* (plant-associated isolates), *P. marginalis* (alfalfa and parsnip pathogens), *P. putida* (rhizosphere and soil isolates), *P. savastanoi* (olive pathogens), *P. stutzeri* (represented by a single clinical isolate), and *P. syringae* (celery, cucumber, tobacco, and tomato isolates or pathogens) (isolates from another eleven *Pseudomonas* spp. were tested, including *P. aeruginosa* PA01 and PA14, and were not found to produce cellulose). For two of the celluloseexpressing isolates, *P. putida* KT2440 and *P. syringae* DC3000, the whole genome sequences were available and SBW25 *wss*-like cellulose synthase operons had been annotated [74-75],

Many environmental pseudomonads can also be induced to form A-L interface biofilms and to express cellulose using WspR19. When expressed *in trans* in wild-type SBW25 it produces the WS phenotype [60, 70], though in other pseudomonads the impact was found to be more variable. In a test of 16 pseudomonads known to form biofilms and express cellulose, WspR19 was found to significantly increase biofilm attachment, strength, and cellulose expression in *P. fluorescens* 54/96, *P. syringae* DC3000, *P. syringae* T1615 and *P. syringae* 6034 [15] (WspR19 induction of cellulose production by SBW25 and DC3000 is shown in Figure 5). WspR19 also induced a WS-like phenotype in *P. putida* KT2440, despite the fact that biofilm-formation or cellulose expression in this pseudomonad had not been observed in the initial survey (cellulose expression was subsequently reported for both wild-type and WspR19-carrying strains under different experimental conditions by [76]). Similarly, nine of ten non-biofilm– forming and non-cellulose expressing *P. syringae* isolates were found to produce biofilms when induced with WspR19, and two of these also expressed detectable levels of cellulose [15]. These findings suggest that biofilm-formation and cellulose expression in pseudomonads closely related to *P. fluorescens* SBW25 are probably controlled by the same c-*di*-GMP– mediated regulatory system or are sensitive to non-specific increases in c-*di*-GMP levels.

Habitat (sample size) A-L interface biofilms Evidence of cellulose

74% 68%

Plant pathogens (n = 57) 6% 26% Plant & soil associated (n = 28) 82% 39% Scottish soil (n = 73)a 95% 76%\* River (n = 57) 82% 5% Indoor & outdoor ponds (n = 50)b 94% 56%

Spoilt cold-stored meat (n = 60)d 77% 28% Mushroom pathogens (n = 26)e 77% 69%

AK & AJS; **b,** B. Varun, AK & AJS; **c,** D.S. Kumar, AK & AJS; **d,** M. Robertson & AJS; and **e,** AK & AJS.

\* Estimated from a sub-sample of 25 isolates. Data compiled from [15] and unpublished research from **a,** R. Ahmed,

**Table 4.** Prevalence of A-L interface biofilm formation and cellulose expression amongst environmental

Pitcher plants (*Sarracenia* spp.) (n = 50)c

pseudomonads

though no experimental reports of either expressing cellulose had been made.

**Figure 7. Activation of the WS phenotype.** The Wrinkly Spreader (WS) phenotype is controlled by the membrane-associated Wsp complex and associated DGC response regulator WspR. **(A)** In wild-type *Pseudomonas fluorescens* SBW25, when an appropriate environmental signal is received the Wsp complex phosphorylates WspR which then results in the production of c-*di*-GMP. However, in the absence of signal, the Wsp complex is silent and c-*di*-GMP levels remain low. **(B)** In the Wrinkly Spreader a mutation in a Wsp subunit (WspF) alters the sensitivity of the Wsp complex such that it activates WspR in the absence of the environmental signal. The resulting increase in c-*di*-GMP activates the membraneassociated cellulose synthase complex to produce cellulose, and also activates the unidentified attachment factor that is also required for the WS phenotype.
