Isolation, screening and phenotypic characterization of biofilm-forming bacteria from farm pond water of semi-arid North Gujarat, India

Introduction

A biofilm is an assemblage of microbial cells irreversibly associated with a surface and enclosed within a self-produced matrix of extracellular polymeric substances (EPS) composed predominantly of polysaccharides, proteins, extracellular DNA and lipids (Flemming et al., 2010) [1] . Biofilms develop on a wide range of living and inert surfaces, from plant roots and animal tissues to water distribution pipes and natural aquatic systems, and their architecture is governed by the nature of the substratum, the hydrodynamics of the system, nutrient availability and grazing pressure. Formation proceeds through a well-described sequence in which planktonic cells attach to a conditioned surface, organise into micro-colonies, and finally encase themselves in a differentiated extracellular matrix from which cells subsequently disperse to colonise new niches (Hall-Stoodley et al., 2004; Davey et al., 2000) [3,2] . The biofilm mode of growth is not merely an architectural curiosity. Cells within a biofilm are markedly more resistant to antimicrobial agents, desiccation and protozoan predation than their planktonic counterparts, and the matrix facilitates horizontal gene transfer and the concentration of secreted metabolites in a defined microenvironment (Costerton et al., 1999; Jefferson, 2004) [4,5] . In soil, these properties underpin the ecological role of biofilms in maintaining nutrient turnover and in sustaining plant growth (Burmølle et al., 2012) [6] . These same properties have prompted a reappraisal of plant growth-promoting rhizobacteria (PGPR). Interactions among PGPR have been explored largely in the planktonic state, yet the behaviour of these organisms as root-associated biofilms is only beginning to be understood (Angus et al., 2013) [7] . Biofilmforming PGPR express plant growth-promoting traits, including diazotrophic nitrogen fixation, indole-3acetic acid synthesis, phosphate and iron solubilisation and ammonia production, more strongly than non-biofilm-forming strains, and they produce substantially higher amounts of antimicrobial compounds (Pandin et al., 2017; Backer et al., 2018; Ajijah et al., 2023) [9,10,8] . Critically, native biofilm-producing PGPR retain their performance under field conditions, where many laboratory-selected planktonic inoculants fail (Timmusk et al., 2005) [11] . Strains isolated for their biofilm-forming ability have since been shown to promote growth in tomato under water deficit (Haque et al., 2020) [12] , in sunflower under salinity (Yasmeen et al., 2020) [13] and in wheat under field conditions (Rafique et al., 2024) [14] . Isolation of such organisms has concentrated on the rhizosphere, but standing water bodies are a rich and comparatively neglected source. Pond water supports diverse attached communities that improve water quality through nitrification, sequester heavy metals and constitute a protein-rich resource in aquaculture systems (Pandey et al., 2014; Kurniawan et al., 2019) [16,15] . Bacteria recovered from such habitats have already been shown to form biofilms and to carry plant growthpromoting attributes (Merina et al., 2011; Nair et al., 2021) [17,18] . Farm ponds in semi-arid regions are of particular interest because their residents experience wide diurnal and seasonal swings in temperature, pH and solute concentration, and are therefore pre-selected for the physiological robustness required of a field-deployed inoculant. Before any such organism can be evaluated agronomically, its identity as a genuine biofilm former must be established by more than one independent method, and its phenotype must be defined. Qualitative tube staining, quantitative microtitre plate assay and Congo red agar morphotyping each detect a different facet of the adherent phenotype, and concordance between them provides considerably stronger evidence than any one assay alone (Christensen et al., 1982; Stepanović et al., 2000; Mathur et al., 2006) [19,20,21] . Equally, the temperature, pH and carbon source at which adherence is maximal define the conditions under which a candidate strain will actually perform. The present study was therefore undertaken to isolate bacteria from the water of two farm ponds in North Gujarat, to identify the strong biofilm formers among them using three independent screening assays, and to characterise the selected isolates morphologically, biochemically and physiologically.

Materials and Methods

Study site and collection of water samples

The investigation was carried out at the Department of Microbiology, Chimanbhai Patel College of Agriculture, Sardarkrushinagar Dantiwada Agricultural University (SDAU), Sardarkrushinagar, Banaskantha district, Gujarat, India. Water was sampled from two farm ponds on the University campus, one at the Vegetable Research Station (24° 19′ 00″ N, 72° 19′ 00″ E) and one at the Centre for Natural Resource Management, CNRM (24° 18′ 43″ N, 72° 16′ 24″ E). The pH of both water bodies ranged from 6.5 to 8.5. Samples were drawn aseptically from the centre of each pond at a depth of 45 cm; 250 mL was collected into a sterile 500 mL conical flask and transported to the laboratory, where it was held at 4 °C until processed.

Isolation of bacteria

A 10 mL aliquot of each water sample was suspended in 90 mL sterile physiological saline (0.85 % NaCl) and agitated on an orbital shaker at 100 rpm and 28 ± 2 °C for 1 h. The suspension was serially diluted to 10 -5 , and 0.1 mL of the 10 -4 and 10 -5 dilutions was spread-plated on nutrient agar and incubated at 28 ± 2 °C for 48 h. Colonies with distinct cultural morphology were picked and purified by repeated streaking on fresh nutrient agar. Pure cultures were maintained on nutrient agar slants at 4 °C.

Screening for biofilm formation

Tube staining assay

Qualitative biofilm formation was assessed following Christensen et al. (1982) [19] . Each isolate (100 µL) was inoculated into 10 mL trypticase soy broth (TSB) supplemented with 5 % sucrose and incubated for 24 h at 37 °C. Tubes were decanted, washed with phosphatebuffered saline (PBS, pH 7.3), dried and stained with 0.1 % crystal violet. Excess stain was removed with deionised water. A visible stained film adhering to the wall and base of the tube was recorded as positive (Mathur et al., 2006) [21] .

Microtitre plate assay

Biofilm biomass was quantified as described by Stepanović et al.

(2000) [20] . Overnight cultures grown statically in TSB with 5 % sucrose at 37 °C were diluted 1:100 in the same medium, and 200 µL was dispensed into the wells of a 96-well polystyrene plate in triplicate. After 24 h of static incubation at 37 °C the contents of each well were removed by gentle tapping, the wells were washed with PBS (pH 7.3) to remove planktonic cells, and adherent biomass was stained with 0.1 % (w/v) crystal violet. Excess stain was washed off with 95 % ethanol and the plates were air-dried. Absorbance was read at 570 nm on a micro-ELISA plate reader. Uninoculated medium served as the negative control and its absorbance defined the cut-off optical density (OD c ). Isolates were classified as non-adherent (OD ≤ OD c ), weakly adherent (𝑂𝐷 c < 𝑂𝐷 ≤ 2 × 𝑂𝐷 c ), moderately adherent (2 × 𝑂𝐷 c < 𝑂𝐷 ≤ 4 × 𝑂𝐷 c ) or strongly adherent (𝑂𝐷 > 4 × 𝑂𝐷 c ).

Congo red agar assay

Isolates were streaked on Mueller-Hinton agar supplemented with 0.8 g L -1 Congo red dye and incubated at 37 °C for 48 h. Black colonies with a dry crystalline consistency were scored as biofilm producers, whereas red to pink colonies were scored as non-producers (Mathur et al., 2006) [21] .

Morphological and cultural characterization

Cell shape and arrangement were recorded after Gram staining following Cappuccino et al. (1992) [22] , using a bright-field compound microscope under oil immersion (E200, Nikon, Japan); images were captured and cell dimensions measured with NIS-Elements Documentation software (Nikon, Japan). The Gram reaction was independently verified by the 3 % KOH string test. A loopful of 18-24 h growth was emulsified in a drop of 3 % KOH on a glass slide for up to 1 min, and the loop was lifted slowly; development of a viscous strand indicated a Gram-negative cell wall, while the absence of stringing indicated a Gram-positive one. Cultural characteristics (colony size, pigmentation, form, elevation and margin) were recorded from well-isolated colonies on nutrient agar after 48 h.

Biochemical characterization

IMViC and carbohydrate utilisation tests were performed with a ready-to-use kit (KB001-10KT, HiMedia, India) according to the manufacturer's protocol. A single colony was inoculated into 5 mL brain heart infusion broth and incubated at 35-37 °C for 4-6 h until turbidity reached ≥ 0.1 OD at 620 nm. Each well was inoculated with 50 µL of the suspension and the plates were incubated at 35-37 °C for 18-24 h, after which Kovac's reagent, methyl red reagent and Barritt's reagents A and B were added to the indole, methyl red and Voges-Proskauer wells respectively. Reactions were read as colour changes per the manufacturer's chart. Catalase activity was assayed by transferring a single colony from a 5-day Luria-Bertani agar plate to a glass slide and adding 2-3 drops of 3 % hydrogen peroxide; immediate effervescence was recorded as positive. Nitrate reduction was tested by inoculating 5 mL sterile nitrate broth and incubating at 25 °C for 8 days, after which two drops each of sulfanilic acid and αnaphthylamine reagents were added to 1 mL of culture; development of a pink to red colour indicated reduction of nitrate to nitrite. Uninoculated broth served as the control.

Extracellular enzyme assays

Amylase and lipase production were assessed by plate assay on substrate agar (Silambarasan et al., 2010; Sheshadari et al., 2011) [24,23] . For amylase, starch agar (meat extract 3 g, soluble starch 2 g, peptic digest of animal tissue 5 g, agar 15 g per litre) was inoculated and incubated at 37 °C for 24-48 h, then flooded with iodine solution; a clear halo around the colony against the blue-black background indicated starch hydrolysis. For lipase, phenol red agar (peptone 5 g, yeast extract 3 g, CaCl 2 1 g, agar 15 g per litre, adjusted to pH 7.4 with 0.1 M NaOH) was autoclaved, cooled to 60 °C and supplemented with 10 mL olive oil and phenol red (1 mg mL -1 ). Plates were incubated at 37 °C for 48 h; a shift of the medium from pink to yellow, reflecting the pH drop caused by liberated fatty acids, indicated lipase activity.

Physiological characterization

The influence of temperature, pH and sucrose concentration on biofilm formation was examined by the microtitre plate assay described in Section 2.3.2, modified as follows. For temperature, 200 µL of diluted culture was dispensed and plates were incubated at 28, 35, 40 and 45 °C for 24 h. For pH, overnight cultures were diluted 1:100 in TSB adjusted to pH 5, 7 or 11 and incubated at 37 °C for 24 h. For carbon source, TSB was supplemented with sucrose at 5, 10, 15 or 20 % and plates were incubated at 37 °C for 24 h. In every case the wells were washed, stained with crystal violet, dried and read at 570 nm as before. Biofilm production was classified as high (OD > 0.500), moderate (OD 0.100-0.500) or poor (OD < 0.100).

Statistical analysis

All assays were performed in triplicate and the results are presented as means. Where treatments were compared, data were subjected to analysis of variance and means were separated by Duncan's New Multiple Range Test (DNMRT) at the 5 % level of significance.

Results

Isolation of bacteria from pond water

Fifteen bacterial colonies with distinct cultural morphology were recovered from the two pond water samples and purified to singlecolony cultures. These were designated sequentially R1 to R15. Isolates R1 to R7 originated from the pond at the Vegetable Research Station and isolates R8 to R15 from the pond at the Centre for Natural Resource Management (Table 1).

Table 1. Bacterial isolates recovered from the two farm pond water samples.

Vegetable Research Station pond

Centre for Natural Resource Management pond

Screening for biofilm formation

All fifteen isolates were screened by the tube staining assay, the microtitre plate assay and the Congo red agar assay. The three methods were fully concordant (Table 2). In the tube assay, five isolates R1, R5, R8, R11 and R12 gave a strongly positive reaction after 24 h, visible as a deep purple, continuous layer of stained biomass adhering firmly to the inner wall and base of the tube, indicating both surface attachment and substantial extracellular matrix production. The remaining ten isolates failed to retain crystal violet after washing and showed neither a film nor a ring along the glass, consistent with an absence of sessile growth under the conditions tested.

The microtitre plate assay placed the same five isolates unambiguously in the strongly adherent class. The negative control gave a stable baseline absorbance of 0.0376, so the thresholds for weak, moderate and strong adherence were 0.038, 0.075 and 0.150 respectively. Isolate R12 gave the highest biofilm biomass (OD 570 0.789), followed by R11 (0.743), R8 (0.719), R5 (0.688) and R1 (0.659); each of these values exceeds the strong-adherence threshold by a factor of more than four (Table 3). On Congo red agar, the same five isolates produced black colonies with a dry, crystalline consistency, the morph type diagnostic of biofilm production, whereas the remaining ten isolates produced smooth red colonies. Because the three independent assays agreed completely, the five positive isolates were retained for detailed characterization and were recoded RBF1 (R1), RBF2 (R5), RBF3 (R8), RBF4 (R11) and RBF5 (R12). [20] relative to the cut-off optical density of the uninoculated control (𝑂𝐷 c = 0.0376).

Morphological and cultural characterization

Microscopic examination after Gram staining showed that all five isolates were rod-shaped and occurred singly rather than in chains or clusters. Isolates RBF1 and RBF3 retained the crystal violet-iodine complex and appeared purple, and were therefore Gram positive, whereas RBF2, RBF4 and RBF5 were decolourised and counterstained pink to red, and were therefore Gram negative. Because Gram staining of environmental isolates can be misleading when decolourisation is imperfectly controlled, the Gram reaction was verified by the 3 % KOH string test. Isolates RBF2, RBF4 and RBF5 produced a viscous strand on lifting the loop, confirming a Gram-negative cell wall, while RBF1 and RBF3 gave no stringing, confirming a Gram-positive cell wall. The two methods agreed for all five isolates (Table 4). On nutrient agar the isolates formed small, compact colonies with dry, often leathery surfaces, giving a conical appearance. Pigmentation ranged from off-white through creamy-white to light yellow. Four of the five isolates (RBF1, RBF2, RBF3 and RBF4) produced colonies of irregular form, whereas RBF5 was circular. Elevation ranged from flat (RBF1) through slightly raised (RBF2) to raised (RBF3, RBF4, RBF5), and colony margins were irregular in RBF1, entire in RBF2 and RBF5, and undulate in RBF3 and RBF4 (Table 5).

In the KOH string test, + denotes formation of a viscous strand (Gram negative) and -denotes no stringing (Gram positive).

Biochemical characterization

The five isolates were differentiated biochemically by the IMViC series, twelve carbohydrate utilisation tests, catalase activity and nitrate reduction (Table 6). Every isolate produced a distinct reaction profile, confirming that the five represent five different organisms rather than replicates of the same strain.

Extracellular enzyme production

After incubation on starch agar and flooding with iodine, isolates RBF1 and RBF2 produced prominent clear zones of hydrolysis surrounding the growth, confirming extracellular amylase activity. In RBF3, RBF4 and RBF5 the dark blue starch-iodine complex extended to the colony margin, indicating no detectable starch degradation.

On phenol red-olive oil agar, none of the five isolates produced a yellow zone of clearance; the medium surrounding every colony remained deep pink, indicating that no fatty acids were liberated and that lipase activity was absent in all five isolates under the conditions tested.

Physiological characterization

Effect of temperature

Biofilm formation increased markedly at 35 °C, which was the optimum for every isolate tested. At this temperature RBF3 gave the highest biofilm biomass (OD 570 0.977), closely followed by RBF2 (0.965), RBF5 (0.880), RBF4 (0.849) and RBF1 (0.840). At 28 °C all five isolates retained a moderate capacity to form biofilm; RBF2 (0.612) and RBF5 (0.596) were the most adaptable at this lower temperature, while RBF1 gave the lowest value (0.485). Raising the temperature to 40 °C reduced biofilm production substantially across the whole set, with values clustering narrowly between 0.455 (RBF4) and 0.520 (RBF2). The constraint became more pronounced at 45 °C, where most isolates reached their lowest recorded yields and RBF4 fell to a minimum of 0.401; RBF3 alone retained a slight tolerance at this temperature (Table 7).

Effect of pH

Biofilm formation was assessed at pH 5, 7 and 11. Neutral conditions were optimal for the majority of the isolates: RBF3, RBF4 and RBF5 all reached their maximum at pH 7, with RBF5 recording the highest overall absorbance of the experiment (OD 570 0.925). Under alkaline conditions biofilm formation was reduced relative to pH 7 but remained appreciable, with RBF3 (0.509) and RBF4 (0.357) retaining a substantial adherent biomass at pH 11. Two isolates departed from this pattern: RBF1 and RBF2 formed most biofilm under mildly acidic conditions, RBF1 peaking at pH 5 with an absorbance of 0.552 (Table 8).

Effect of sucrose concentration

All five isolates formed biofilm at every sucrose concentration tested, and in each case production was maximal at 10 % sucrose. RBF2 maintained a strong adherent phenotype across the whole range, peaking at 10 % with an absorbance of 0.970 and retaining high biofilm biomass at 15 % (0.840) and 20 % (0.759). The remaining four isolates (RBF1, RBF3, RBF4 and RBF5) showed weak to moderate formation, characterised by a steady rise from 5 % to 10 % sucrose followed by a sharp decline at 15 % and 20 % (Table 9).

Discussion

Five of the fifteen bacteria recovered from two semi-arid farm ponds, a recovery rate of one in three, were strong biofilm formers. This proportion is comparable with that reported for other aquatic habitats: Nair et al. (2021) [18] found that seven of twelve isolates from a water treatment tank formed biofilm in the tube assay within 24 h, and Merina et al. (2011) [17] recovered biofilm-forming bacilli from a marine seagrass habitat. Standing farm water therefore appears to be a productive source of adherent bacteria, and one that is logistically far simpler to sample than the rhizosphere. The principal methodological strength of this screening is that three assays with different readouts agreed completely. The tube assay detects gross adherence and matrix retention but is qualitative and prone to observer bias; the microtitre plate assay quantifies attached biomass but can register non-specific staining; and the Congo red agar assay detects the dry, rugose morphotype associated with exopolysaccharide and amyloid fibre production, a phenotype known to be regulated independently of attachment itself (Römling, 2005; Milanov et al., 2015) [26,25] . Individual assays are known to disagree, and Mathur et al. (2006) [21] concluded from a direct comparison of the three that agreement should not be assumed. The complete concordance observed here, across fifteen isolates rather than five, gives considerably greater confidence that RBF1 to RBF5 are genuine biofilm formers than any single assay could. The margin was also large: all five gave OD 570 values more than four times the cut-off, with the lowest (0.659) more than seventeen times the control absorbance, so the classification does not depend on where the boundary between moderate and strong adherence is drawn. Phenotypically the five isolates were uniform in some respects and diverse in others. All were single, rod-shaped cells and all were catalase positive. The rod morphology is consistent with the dominance of Bacillaceae, Paenibacillaceae and Gram-negative Proteobacteria among reported biofilm-forming PGPR (Bais et al., 2004; Timmusk et al., 2005; Ansari et al., 2019) [28,11,27] . Universal catalase activity is not incidental. Catalase neutralises hydrogen peroxide and so protects DNA, lipids and proteins from oxidative damage; its presence in all five isolates is consistent with the observation that biofilm-forming bacteria generally carry stronger antioxidant defences than planktonic strains, a trait that supports persistence on root surfaces and under the salinity, drought and oxidative stress encountered in the field (Kloepper et al., 1989; Flemming et al., 2010) [29,1] . Yasmeen et al. (2020) [13] likewise recorded catalase activity in all their biofilmforming rhizobacteria and linked it directly to enhanced salt tolerance in sunflower. The decision to verify the Gram reaction by an independent chemical method deserves emphasis. Gram staining depends on controlled decolourisation, and certain spore-forming genera, notably Bacillus and Clostridium, are known to stain Gram-negative despite possessing a Gram-positive cell wall. The 3 % KOH string test, which depends on the differential lysis of the Gram-negative envelope and the consequent release of viscous chromosomal DNA, supplied an orthogonal check. That the two methods agreed for all five isolates means that the assignment of RBF1 and RBF3 as Gram positive and RBF2, RBF4 and RBF5 as Gram negative is secure, and that any subsequent genomic DNA extraction protocol can be selected on a sound basis. Colony morphology on nutrient agar small, compact colonies with dry, leathery, often conical surfaces is itself informative, since a dry rugose surface reflects abundant extracellular matrix and correlates with the black, crystalline morph type on congo red agar. The range of forms, margins and pigments recorded here is consistent with that described by Hussain et al. (2013) [30] for bacteria isolated from water sources, who reported round to irregular forms, smooth to dry surfaces, and whitish, cream and yellow pigmentation in a comparable collection. The biochemical profiles distinguished all five isolates from one another. Universal citrate utilisation combined with an inability to use adonitol, arabinose, lactose or sorbitol indicates organisms adapted to simple rather than complex carbon sources, which is consistent with a nutrient-limited pond water origin. The isolates differed in indole, methyl red and Voges-Proskauer reactions and in their use of glucose, mannitol, rhamnose and sucrose, and this combination of characters is the conventional basis for provisional generic placement (Sujana et al., 2013; Sonkusale et al., 2015) [32,31] . Nitrate reduction in RBF1, RBF2 and RBF5 is of particular agronomic interest, since nitrate-reducing biofilms contribute to nitrogen cycling in both aquatic and rhizosphere environments. Yasmeen et al. (2020) [13] similarly used nitrate reductase activity and citrate utilisation to separate their biofilmforming isolates. Extracellular enzyme production was more restricted than in some published collections. Amylase was confined to RBF1 and RBF2, and lipase was absent throughout. Nair et al. (2021) [18] found that all seven of their biofilm-forming strains produced both amylase and lipase, and suggested that exoenzymes assist adherence to solid surfaces. The present results indicate that neither enzyme is required for strong adherence, since RBF3, RBF4 and RBF5 produced the highest microtitre plate absorbances of the set while producing neither enzyme. Attachment in these isolates is therefore likely to depend on matrix polysaccharide and surface adhesins rather than on substrate hydrolysis. The practical implication is positive: a strain that does not secrete lipase is less likely to degrade seed coat lipids during seed treatment. The physiological data define the operating window within which these organisms will perform. A temperature optimum of 35 °C in all five isolates, with substantial biofilm still formed at 28 °C and measurable biofilm retained at 40 and 45 °C, matches the thermal regime of North Gujarat soils during the kharif and summer seasons and indicates that adherence will not collapse under field heat. Sonkusale et al. (2015) [31] reported an analogous pattern, with maximum biofilm at 37 °C, retention at 50 °C in some isolates, and minimal formation at 25 °C. The pH response was more revealing. That RBF3, RBF4 and RBF5 peaked at pH 7 accords with the general finding that neutral conditions favour cell attachment and EPS matrix production (Sonkusale et al., 2015; Cai et al., 2013) [31,33] . More interesting is the retention of appreciable biofilm at pH 11 by RBF3 and RBF4, and the departure of RBF1 and RBF2, which formed most biofilm at pH 5. Alkaline calcareous soils predominate across much of North Gujarat, so alkali-tolerant adherence is a directly useful trait, while acid-tolerant isolates broaden the range of soils in which the collection as a whole could be deployed. Taken together the five isolates cover a pH span from 5 to 11, which is unusually wide for a set of this size. Sucrose supported biofilm formation in all isolates with a consistent optimum at 10 %, above which formation declined sharply in four of the five. The decline is most plausibly osmotic: at 15 and 20 % sucrose the water activity of the medium falls far enough to impose a solute stress that outweighs the benefit of additional carbon for matrix synthesis. RBF2 was the exception, maintaining absorbances of 0.840 and 0.759 at 15 and 20 % respectively, which points to superior os motolerance. Since osmotic and salt stress tolerance is one of the central arguments for using biofilm-forming inoculants in the first place (Qurashi et al., 2012; Ansari et al., 2019; Ajijah et al., 2023) [34,27,8] , RBF2 merits particular attention in any subsequent evaluation under saline or drought conditions. The wider relevance of this collection lies in the accumulating evidence that biofilm-forming rhizobacteria outperform planktonic inoculants where it matters. Biofilm-forming strains express plant growthpromoting traits more strongly, produce more antimicrobial metabolites and survive better in competitive soil (Pandin et al., 2017; Backer et al., 2018) [9,10] , and they maintain performance under field conditions where laboratory-selected planktonic strains commonly fail (Timmusk et al., 2005 [11,12,13] have each demonstrated growth promotion by biofilm-forming isolates in tomato, sunflower and wheat respectively. The five isolates reported here have been defined phenotypically to a standard sufficient to support that next step. Two limitations should be stated. First, the isolates are characterised phenotypically and have not been assigned to species here; molecular identification based on 16S rRNA gene sequencing is required before any taxonomic claim can be made, and is reported separately. Second, biofilm screening was conducted at 37 °C on a sucrose-rich laboratory medium, conditions that optimise the assay but do not reproduce the rhizosphere; adherence on root surfaces in situ remains to be demonstrated.

Conclusion

Fifteen bacteria were isolated from the water of two farm ponds in semi-arid North Gujarat, and five of them recoded RBF1 to RBF5 were identified as strong biofilm formers by three independent and fully concordant assays. The five isolates are rod-shaped, singlecelled and uniformly catalase positive; two are Gram positive and three Gram negative, as confirmed by both Gram staining and the 3 % KOH string test. Each has a distinct biochemical profile, all utilise citrate, three reduce nitrate, two produce extracellular amylase and none produces lipase. Biofilm formation is maximal at 35 °C and 10 % sucrose in every isolate, and the set collectively spans pH 5 to pH 11, with RBF2 additionally tolerating sucrose concentrations up to 20 %. Farm pond water is thus confirmed as a convenient and productive source of robust biofilm-forming bacteria, and RBF1 to RBF5 constitute a phenotypically defined collection suitable for molecular identification and for evaluation as biofilm-based bioinoculants.

Acknowledgement

AcknowledgementsThe authors thank the Department of Microbiology, Chimanbhai Patel College of Agriculture, Sardarkrushinagar Dantiwada Agricultural University, for laboratory facilities. [Add funding and any further acknowledgements.]

References

1. Flemming HC, Wingender J. The biofilm matrix. Nature Reviews Microbiology. 2010;8:623-633.
2. Davey ME, O'Toole GA. Microbial biofilms: from ecology to molecular genetics. Microbiology and Molecular Biology Reviews. 2000;64:847-867.
3. Hall-Stoodley L, Costerton JW, Stoodley P. Bacterial biofilms: from the natural environment to infectious diseases. Nature Reviews Microbiology. 2004;2:95-108.
4. Costerton JW, Stewart PS, Greenberg EP. Bacterial biofilms: a common cause of persistent infections. Science. 1999;284:13181322.
5. Jefferson KK. What drives bacteria to produce a biofilm? FEMS Microbiology Letters. 2004;236:163-173.
6. Burmølle M, Kjøller A, Sørensen SJ. An invisible workforce: biofilms in the soil. In: Lear G, Lewis G, editors. Microbial Biofilms -Current Research and Applications. Norfolk: Caister Academic Press; 2012:61-72.
7. Angus AA, Hirsch AM. Biofilm formation in the rhizosphere: multispecies interactions and implications for plant growth. In: Molecular Microbial Ecology of the Rhizosphere. 2013:701-712.
8. Ajijah N, Fiodor A, Pandey A, Rana A, Pranaw K. Plant growthpromoting bacteria (PGPB) with biofilm-forming ability: a multifaceted agent for sustainable agriculture. Diversity. 2023;15:112.
9. Pandin C, Le Coq D, Canette A, Aymerich S, Briandet R. Should the biofilm mode of life be taken into consideration for microbial biocontrol agents? Microbial Biotechnology. 2017;10:719-734.
10. Backer R, Rokem JS, Ilangumaran G, Lamont J, Praslickova D, Ricci E, Subramanian S, Smith DL. Plant growth-promoting rhizobacteria: context, mechanisms of action, and roadmap to commercialization of biostimulants for sustainable agriculture. Frontiers in Plant Science. 2018;9:1473.
11. Timmusk S, Grantcharova N, Wagner EGH. Paenibacillus polymyxa invades plant roots and forms biofilms. Applied and Environmental Microbiology. 2005;71:7292-7300.
12. Haque MM, Mosharaf MK, Khatun M, Haque MA, Biswas MS, Islam MS, Siddiquee MA. Biofilm-producing rhizobacteria with multiple plant growth-promoting traits promote growth of tomato under water-deficit stress. Frontiers in Microbiology. 2020;11:542053.
13. Yasmeen T, Ahmad A, Arif MS, Mubin M, Rehman K, Shahzad SM, Wijaya L. Biofilm-forming rhizobacteria enhance growth and salt tolerance in sunflower plants by stimulating antioxidant enzyme activity. Plant Physiology and Biochemistry. 2020;156:242-256.
14. Rafique M, Naveed M, Mumtaz MZ, Niaz A, Alamri S, Siddiqui MH, Mustafa A. Unlocking the potential of biofilm-forming plant growth-promoting rhizobacteria for growth and yield enhancement in wheat (Triticum aestivum L.). Scientific Reports. 2024;14:15546.
15. Kurniawan A, Salamah LN, Susanti YAD, Damaika MA, Guspinta YRP, Lestari WA, Ulfa SM. Analysis of biofilms as biomonitoring agent for Cu 2+ and Pb 2+ pollution in the lotic ecosystem. IOP Conference Series: Earth and Environmental Science. 2019;239:012026.
16. Pandey PK, Bharti V, Kumar K. Biofilm in aquaculture production. African Journal of Microbiology Research. 2014;8:14341443.
17. Merina M, Lipton AP, Wesley SG. Isolation, characterization and growth of biofilm-forming bacteria Bacillus pumilus from the seagrass Halodule pinifolia of Kanyakumari coast. Indian Journal of Marine Sciences. 2011;40:443-448.
18. Nair S, Kwatra N, Abraham J. Isolation and characterization of biofilm-producing bacteria from water treatment tank. Current Trends in Biotechnology and Pharmacy. 2021;15:34.
19. Christensen GD, Simpson WA, Bisno AL, Beachey EH. Adherence of slime-producing strains of Staphylococcus epidermidis to smooth surfaces. Infection and Immunity. 1982;37:318326. International Journal of Biology Sciences https://www.biologyjournal.net 26649934, 2026, 8(9), Downloaded from https://www.doi.org/10.33545/26649926.2026.v8.i9.b.996 by International Journal of Biology Sciences on [25/09/2026]. See the Terms and Conditions (https://www.biologyjournal.net) for rules of use; OA articles are governed by the applicable Creative Commons License
20. Stepanović S, Vuković D, Dakić I, Savić B, Švabić-Vlahović M. A modified microtitre-plate test for quantification of staphylococcal biofilm formation. Journal of Microbiological Methods. 2000;40:175-179.
21. Mathur T, Singhal S, Khan S, Upadhyay UP, Fatma F, Rattan A. Detection of biofilm formation among the clinical isolates of staphylococci: an evaluation of three different screening methods. Indian Journal of Medical Microbiology. 2006;24:25-29.
22. Cappuccino JG, Sherman N. Microbiology: A Laboratory Manual, 3rd ed. Redwood City: Benjamin/Cummings; 1992:125179.
23. Sheshadari P, Abraham J. A study on Enterococcus faecalis antibiotic resistance pattern isolated from patients with urinary tract infection. International Journal of Universal Pharmaceutical and Life Sciences. 2011;1:129-133.
24. Silambarasan S, Prasad VH, Balaji R, Abraham J. A study on antimicrobial property of salt-tolerant bacteria isolated from Indian mangrove forest. Report and Opinion. 2010;2:27-32.
25. Milanov DS, Prunić BZ, Velhner MJ, Pajić ML, Čabarkapa IS. rdar morphotype -a resting stage of some Enterobacteriaceae. Food and Feed Research. 2015;42:43-50.
26. Römling U. Characterization of the rdar morphotype, a multicellular behaviour in Enterobacteriaceae. Cellular and Molecular Life Sciences. 2005;62:1234-1246.
27. Ansari FA, Ahmad I. Isolation, functional characterization and efficacy of biofilm-forming rhizobacteria under abiotic stress conditions. Antonie van Leeuwenhoek. 2019;112:1827-1839.
28. surfactin production. Plant Physiology. 2004;134:307-319.
29. Kloepper JW, Lifshitz R, Zablotowicz RM. Free-living bacterial inocula for enhancing crop productivity. Trends in Biotechnology. 1989;7:39-44.
30. Hussain T, Roohi A, Munir S, Ahmed I, Khan J, Edel-Hermann V, Anees M. Biochemical characterization and identification of bacterial strains isolated from drinking water sources of Kohat, Pakistan. African Journal of Microbiology Research. 2013;7:1579-1590.
31. Sonkusale KD, Tale VS. Isolation and characterization of biofilmforming bacteria from oral microflora. International Journal of Current Microbiology and Applied Sciences. 2015;4:118-127.
32. Sujana K, Abraham KP, Mikkili I, Prabhakar KV. Biochemical and molecular characterization of biofilm-producing bacteria. International Journal of Pharma and Bio Sciences. 2013;4:702712.
33. Cai W, De La Fuente L, Arias CR. Biofilm formation by the fish pathogen Flavobacterium columnare: development and parameters affecting surface attachment. Applied and Environmental Microbiology. 2013;79:5633-5642.
34. Qurashi AW, Sabri AN. Bacterial exopolysaccharide and biofilm formation stimulate chickpea growth and soil aggregation under salt stress. Brazilian Journal of Microbiology. 2012;43:11831191. How to cite this article: Parmar RB, Singh N, Patel MM, Gangwar G, Yadav A, Tapre P, et al. Isolation, screening and phenotypic characterization of biofilm-forming bacteria from farm pond water of semi-arid North Gujarat, India. International Journal of Biology Sciences. 2026;8(9):110-117. doi: https://ww