Qualitative and quantitative evaluation of biofilm formation in indigenous Bacillus thuringiensis isolates

Introduction

Bacillus thuringiensis (Bt) is a facultative anaerobic, gram-positive, motile, spore-forming, rod-shaped bacterium (Martin and Travers., 1989) [10] widely used in biological pest control due to its production of insecticidal crystal proteins (Cry and Cyt toxins) that target specific insect pests, particularly those in the orders Lepidoptera, Diptera, and Coleoptera (Vidyarthi et al., 2002) [19] . These proteins are known for their high specificity and environmental safety, making Bt one of the most successful microbial biopesticides in the world. Bt accounts for over 90% of the global biopesticide market and is considered the most successful microbial pesticide used in agriculture, forestry, and public health (Sanchis and Bourguet., 2008) [15] . Beyond toxin production, Bt has the ability to form biofilms (Nay et al., 2016). Bacterial biofilms are aggregates of bacteria embedded in extracellular polymeric substances (EPSs) produced by the bacteria themselves. The EPS matrix typically includes extracellular polysaccharides, proteins, lipids, and DNA, enabling the biofilm to adhere to a range of interfaces such as solid-liquid, solid-gas, liquid-liquid, and liquid-gas surfaces (Flemming et al., 2016 [6] ; Flemming and Wuertz., 2019) [5] . Bacteria are capable of colonizing nearly all natural and synthetic surfaces by forming biofilms (Hall-Stoodley et al., 2004 [7] ; Sweet et al., 2011 [18] ; Flemming and Wuertz, 2019) [5] . Compared with planktonic cells, the most important feature of bacterial biofilm is that it can help bacteria resist different environmental stresses such as ultraviolet (UV) radiation, antibiotic treatment, drying, temperature variation, oxidative stress, nutrient depletion, and pH change (Rinaudi and Giordano, 2010 [13] ; Bernbom et al., 2011 [2] ; Koerdt et al., 2011 [9] ; Dang and Lovell, 2016) [3] . Many B. thuringiensis strains can form biofilms that float on the surface of liquid culture media (Wijman et al., 2007 [20] ; Auger et al., 2009) [1,16] . These surface biofilms have a distinct structure, typically featuring a dense ring that adheres to the vessel wall and encircles the floating pellicle, which often exhibits visible protrusions (Fagerlund et al., 2014) [4] . Conventional culturing of Bt typically involves shaking/incubation under aerated conditions. However, this requires costly infrastructure and high energy inputs (Rowe and Margaritis, 2004) [14] . Static culture, on the other hand, can simulate simple conditions suitable for farmer-level or resource-constrained applications. Such non-agitated systems are more accessible and can serve as farmer-friendly methods for microbial mass culturing. The current study aims to screen indigenous Bt isolates for their biofilm production ability under static (non-agitated) conditions. Wet pellet weight was used as a direct indicator of biofilm formation, and OD₆₀₀ readings were recorded to observe growth dynamics under static incubation. This approach can help identify robust, locally adapted Bt strains for further development as biocontrol agents.

Materials and Methods

Bacterial Isolates

A total of twenty indigenous Bacillus thuringiensis (Bt) isolates collected from various agro-ecological zones of Maharashtra and maintained at the Department of Entomology and Biotechnology centre, Post Graduate Institute, Dr. PDKV, Akola, were used in this study. The standard strain Bacillus thuringiensis subsp. kurstaki HD-1 was included for comparison. The details of Bt isolates, used for evaluation of biofilm production are presented in (Table 1). The previously isolated Bt cultures were sub-cultured and maintained on nutrient agar plates at 4 °C for further use. [4] . Observations on biofilm formation were recorded at 24, 48, 72, and 96 hours post-inoculation.

Qualitative Assessment of Biofilm Formation

At each time point (24 h, 48 h, 72 h, and 96 h), the test tubes were visually examined for the presence of ring (formed at the liquid-air interface) and pellicle (floating biofilm) structures. Based on the coverage, density, and strength of the biofilm, isolates were categorized as:

Quantitative Estimation of Biofilm Formation

After 96 hours, both the ring and the pellicle part of biofilm were harvested separately into sterile microcentrifuge tubes (Mengmeng et al., 2021). For ring recovery, the biofilm was manually scrapped from the sides of the tubes using a micropipette tip and transferred into microcentrifuge tube and the floating pellicle at the bottom was recovered using a micropipette and transferred to a separate microcentrifuge tube (Auger et al., 2005). The samples were then centrifuged at 8000 rpm for 5 minutes, and the resulting pellet yield was recorded using a digital precision balance and their corresponding OD 600 values were determined using spectrophotometer.

Statistical Analysis

All experiments were performed in triplicate, and the results were expressed as mean±standard error (SE). Differences in biofilm formation among the Bacillus thuringiensis isolates were analysed using one-way Analysis of Variance (ANOVA). The statistical analysis was conducted using GraphPad Prism version 9.0. A p-value less than 0.05 (p<0.05) was considered statistically significant.

Results and Discussion

Qualitative Assessment of Biofilm Formation

Biofilm development by Bacillus thuringiensis (Bt) isolates was visually evaluated through the observation of pellicle and ring structures in static culture tubes at different incubation intervals (24, 48, 72, and 96 hours). At 24 hours, isolate SA-18 alone exhibited ring formation. At 48 hours, isolates SY-4, SJa-2, SA-22, SDa-1, and the standard strain HD-1 produced both ring and pellicle structures. Three isolates SGd-1, SA-18, and SUd-2 formed only ring, while 12 isolates viz., (SA-6, SA-20, SAk-6, SAk-9, SGn-5, SGn-4, SJa-1, SHi-1, SDa-2, SMa-1, SAk-12 and SUd-1) exhibited only pellicle formation. Whereas, SBn-2 showed no visible biofilm. At 72 hours, 18 isolates viz., (SA-6, SA-20, SGd-1, SA-18, SAk-6, SAk-9, SGn-5, SY-4, SUd-2, SJa-2, SJa-1, SHi-1, SDa-2, SDa-1, SA-22, SMa-1, SUd-1) including HD-1, produced both ring and pellicle structures, while three isolates (SBn-2, SGn-4, and SAk-12) produced only pellicle. At 96 hours, the same three isolates (SBn-2, SGn-4, and SAk-12) continued to produce only the pellicle part, while the remaining eighteen isolates consistently developed both ring and pellicle biofilms.

Based on the intensity and coverage of biofilm formation, eight isolates (HD-1, SGd-1, SA-18, SAk-6, SAk-9, SY-4, SJa-2, and SA-22) were categorized as strong ring biofilm producers, while ten isolates (SA-6, SA-20, SGn-5, SUd-2, SJa-1, SHi-1, SDa-2, SDa-1, SMa-1, and SUd-1) exhibited weak ring formation. With respect to pellicle development, fifteen isolates (HD-1, SA-6, SA-20, SGd-1, SAk-6, SAk-9, SY-4, SUd-2, SHi-1, SDa-2, SDa-1, SA-22, SAk-12, and SUd-1) demonstrated strong pellicle formation, while six isolates (SA-18, SGn-5, SBn-2, SGn-4, SJa-1, SJa-2, and SMa-1) were identified as weak pellicle producers (Table 2; Figure 1). The results obtained in the present study regarding the variability in biofilm formation among different Bacillus thuringiensis isolates over time are in agreement with earlier findings. Karina et al. (2014) [8] and Mengmeng et al. (2021) also reported considerable heterogeneity in the biofilmforming ability among various Bt isolates. Similarly, Xingchen Zhao et al. (2024) [21] observed that the Bt strain SA-11 exhibited strong biofilm formation, whereas ABTS-351 showed weak to moderate formation, and ABTS-1857 failed to form biofilm altogether. Nay El-Khoury et al. (2016) [12] studied biofilm development in Bt strains over time intervals from 16 to 96 hours and observed distinct structures ring and pellicle during the progression. According to their findings, no visible ring or pellicle was observed at 16 hours. By 20 hours, a thin pellicle surrounded by a ring began to appear, and at 24 hours, the biofilm developed a well-defined architecture characterized by a thick ring and dense pellicle layer. With prolonged incubation (36, 48, and 72 hours), both ring and pellicle structures became more prominent and welldeveloped, indicating dynamic changes in biofilm architecture over time. These observations align with our findings, where ring and pellicle formation varied significantly among isolates and intensified with longer incubation durations.

Quantitative Estimation of Biofilm Formation

To evaluate the biofilm yield potential of Bacillus thuringiensis (Bt) isolates, the ring and pellicle components were harvested separately following 96 hours of static incubation. The biofilm materials were gently collected using a sterile pipette and centrifuged at 8000 rpm for 5 minutes to obtain biomass pellets. Quantitative assessment included measuring the wet weight of the biomass (pellet weight) and the optical density at 600 nm (OD₆₀₀) to estimate cell density.

For each isolate, two sets of parameters were recorded: (i) Ring portion-pellet weight and OD₆₀₀ value, and (ii) Pellicle portion-pellet weight and OD₆₀₀ value.

Yield Assessment of Bt Isolates in Ring Biofilm

Ring Biofilm Pellet Biomass

The ring-associated biomass (Table 3; Figure 2) exhibited considerable variation among the tested Bacillus thuringiensis (Bt) isolates. Among the 18 isolates that produced visible ring structures, the pellet biomass ranged from 11.50±1.50 mg to 60.50±27.50 mg. The standard strain, HD-1, recorded a ring biomass of 26.00±12.00 mg. Notably, the indigenous isolate SA-18 yielded the highest biomass (60.50±27.50 mg), which was approximately 2.33 times greater than that of HD-1. This was followed by SJa-2 (51.50±23.50 mg, 1.98 times) and SY-4 (35.50±10.50 mg, 1.36 times). Isolates such as SGd-1 (25.00±8.00 mg), SGn-5 (24.50±0.50 mg), and SAk-6 (24.00±1.00 mg) produced ring biomass comparable to that of the standard strain. Conversely, isolates including SGn-4, SBn-2, and SAk-12 exhibited no measurable ring biomass (0.00±0.00 mg), suggesting poor or absent growth in ring form. Statistical analysis revealed a significant difference among the treatments (p < 0.05).

Ring Biofilm OD₆₀₀ Analysis

The optical density (OD₆₀₀) values for the ring biomass of Bt isolates, as presented in Table 3 and Figure 3, ranged from 0.00 to 2.28. The standard strain HD-1 recorded an OD₆₀₀ of 1.97. Among the indigenous isolates, SA-18 (2.28), SJa-2

(2.02), and SY-4 (1.98) recorded higher OD₆₀₀ values than HD-1, indicating a greater cell density, which corresponded well with their respective ring pellet biomasses. Isolates such as SGd-1 (1.93) and SGn-5 (1.92) showed OD₆₀₀ values comparable to the standard strain, suggesting similar biofilmforming ability. In contrast, isolates like SA-22 (0.54), SMa-1 (0.62), SGn-4 (0.00), SBn-2 (0.00), and SAk-12 (0.00) exhibited markedly lower or negligible OD₆₀₀ values, indicative of weak or absent ring biofilm development under static conditions. The quantitative assessment of pellicle biomass among the Bacillus thuringiensis isolates exhibited marked variability, as presented in Table 4 and Figure 2. The pellicle pellet weight ranged from 25.50±21.00 mg to 113.50±1.00 mg across the tested isolates. The reference strain, HD-1, yielded a pellicle biomass of 105.00±3.00 mg. Among the indigenous isolates, SAk-12 recorded the highest pellicle biomass (113.50±1.00 mg), which was 1.08 times greater than the standard strain. This was followed by SAk-6 (106.50±2.00 mg, 1.01 times). Other isolates such as SAk-9 (91.00±4.00 mg), SHi-1 (76.00±5.00 mg), and SGd-1 (73.50±6.00 mg) also demonstrated substantial biomass production. In contrast, isolates including SA-18 (25.50±21.00 mg) and SJa-2 (40.50±2.00 mg) produced significantly lower pellicle biomass, suggesting a weaker capacity for pellicle development. Statistical analysis confirmed that the differences observed among the treatments were statistically significant (p < 0.05), indicating true variability in pellicle-forming ability across the Bt isolates.

Pellicle Biofilm OD₆₀₀ Analysis

The OD₆₀₀ values measured for the pellicle component of Bacillus thuringiensis isolates demonstrated considerable variation, ranging from 0.34 to 3.95 (Table 4; Figure 3). The standard strain HD-1 recorded an OD₆₀₀ of 3.02, serving as a benchmark for comparison. Notably, SAk-12 (3.95) and SAk-6 (3.22) surpassed the standard strain, indicating higher cell density and more robust pellicle formation. These were followed by isolates SAk-9 (2.85) and SHi-1 (2.62), which also recorded strong growth profiles. In contrast, isolates SA-18 (0.34) and SJa-2 (0.43) exhibited markedly lower OD₆₀₀ values in the pellicle phase. Interestingly, despite their reduced pellicle density, these isolates had earlier demonstrated significant biomass accumulation in the ring component, suggesting a preferential biofilm formation near the liquid-air interface rather than in the submerged pellicle layer. The present study revealed significant variability in biomass production among Bacillus thuringiensis (Bt) isolates, both in the ring and pellicle phases. Certain isolates showed high biomass accumulation in one form but not the other, underscoring strain-specific growth patterns likely influenced by physiological and environmental factors Sonia Paytubi et al., 2017 [17] . A strong positive correlation was observed between OD₆₀₀ values and wet pellet weights across both biomass phases, validating OD₆₀₀ as a reliable indicator of biomass yield. High-performing isolates such as PDKV-Bt-SAk-6, PDKV-Bt-SAk-9, and PDKV-Bt-SGd-1 consistently recorded elevated OD₆₀₀ values along with substantial biomass in both the ring and pellicle components. Conversely, PDKV-Bt-SAk-12 exhibited negligible ring formation yet demonstrated the highest pellicle OD₆₀₀ and biomass, suggesting a preference for bottom-associated growth, likely governed by isolate-specific traits. The standard strain HD-1 maintained balanced growth across both components, supporting its continued use as a reference standard. The observed differences in biomass distribution patterns between ring and pellicle phases highlight the diverse biofilm-forming abilities of Bt isolates. These findings are consistent with earlier studies. Nay El-Khoury et al. (2016) [12] reported a proportional relationship between OD₆₀₀ and biomass (spores and vegetative cells), with pellicle OD₆₀₀ values approximately 1. [8] further demonstrated OD₆₂₀ as a reliable metric for biofilm quantification in 40 environmental Bt strains, categorizing several strains as strong biofilm producers based on their OD values. Overall, integrating OD₆₀₀ analysis with biomass measurements offers a comprehensive understanding of growth behaviour and provides a practical approach for selecting high-yielding Bt strains for biocontrol and industrial applications.

Acknowledgement

AcknowledgementsThis study is a part of thesis research of first author submitted to Department of Entomology, Post Graduate Institute, Dr. PDKV Akola. The authors are grateful to Department of Entomology, Post Graduate Institute, Dr. PDKV, Akola, MS, India for providing all the necessary facilities for conducting the present research work. Authors are also grateful to the student's advisory committee members for their critical comments and valuable suggestions during the study. International Journal of Advanced Biochemistry Research https://www.biochemjournal.com

References

1. Auger S, Ramarao N, Faille C, Fouet A, Aymerich S, Gohar M. Biofilm formation and cell surface properties among pathogenic and nonpathogenic strains of the Bacillus cereus group. Appl Environ Microbiol. 2009;75:6616-6618.
2. Bernbom N, Vogel BF, Gram L. Listeria monocytogenes survival of UV-C radiation is enhanced by presence of sodium chloride, organic food material and by bacterial biofilm formation. Int J Food Microbiol. 2011;147:6973.
3. Dang H, Lovell C. Microbial surface colonization and biofilm development in marine environments. Microb Mol Biol Rev. 2016;80:91-138.
4. Fagerlund A, Dubois T, Okstad OA, Verplaetse E, Gilois N, Bennaceur I, et al. SinR controls enterotoxin expression in Bacillus thuringiensis biofilms. PLoS ONE. 2014;9:e87532.
5. Flemming HC, Wuertz S. Bacteria and archaea on Earth and their abundance in biofilms. Nat Rev Microbiol. 2019;17:247-260.
6. Flemming HC, Wingender J, Szewzyk U, Steinberg P, Rice SA, Kjelleberg S. Biofilms: An emergent form of bacterial life. Nat Rev Microbiol. 2016;14:563-575.
7. Hall-Stoodley, Costerton JW, Stoodley P. Bacterial biofilms: from the natural environment to infectious diseases. Nat Rev Microbiol. 2004;2:95-108.
8. Karina Garcia J, Bravo A, Diaz J, Gutierrez D, Torres PV, Gomez de Leon P. Variability of Bacillus thuringiensis strains by ERIC-PCR and biofilm formation. 2014;70:10-18.
9. Koerdt A, Orell A, Trong Khoa P, Mukherjee J, Wlodkowski A, Karunakaran E. Macromolecular fingerprinting of Sulfolobus species in biofilm: A transcriptomic and proteomic approach combined with spectroscopic analysis. J Proteome Res. 2011;10:41054119.
10. Martin PAW, Travers RS. Worldwide abundance and distribution of Bacillus thuringiensis isolates. Appl Environ Microbiol. 1989;55:2437-2442.
11. Li M, Shu C, Ke W, Li X, Yu Y, Guan X, et al. Plant Polysaccharides Modulate Biofilm Formation and Insecticidal Activities of Bacillus thuringiensis Strains. Front Microbiol. 2021;12.
12. El-Khoury N, Majed R, Perchat S, Kallassy M, Lereclus D, Gohar M. Spatio-Temporal Evolution of Sporulation in Bacillus thuringiensis Biofilm. Front Microbiol. 2016;7.
13. Rinaudi LV, Giordano W. An integrated view of biofilm formation in rhizobia. FEMS Microbiol Lett. 2010;304:1-11.
14. Rowe G, Margaritis A. Bioprocess design and economic analysis for the commercial production of environmentally friendly bioinsecticides from Bacillus thuringiensis HD-1 kurstaki. Biotechnol Bioeng. 2004;86(4):377-384.
15. Sanchis V, Bourget D. Bacillus thuringiensis: applications in agriculture and insect resistance management. Agron Sustain Dev. 2008;28:11-20.
16. Auger S, Ramarao N, Faille C, Fouet A, Aymerich S, Gohar M. Biofilm Formation and Cell Surface Properties among Pathogenic and Nonpathogenic Strains of the Bacillus cereus Group. Appl Environ Microbiol. 2009;75(20):6616-6618.
17. Paytubi S, Cansado C, Madrid C, Balsalobre C. Nutrient Composition Promotes Switching between Pellicle and Bottom Biofilm in Salmonella. Front Microbiol. 2017;8.
18. Sweet MJ, Croquer A, Bythell JC. Development of bacterial biofilms on artificial corals in comparison to surface-associated microbes of hard corals. PLoS One. 2011;6:e21195.
19. Vidyarthi AS, Tyagi RD, Valero JR, Surampalli RY. Studies on the production of B. thuringiensis based biopesticides using wastewater sludge as a raw material. Water Res. 2002;36:4850-4860.
20. Wijman JG, De Leeuw PP, Moezelaar R, Zwietering MH, Abee T. Air-liquid interface biofilms of Bacillus cereus: formation, sporulation, and dispersion. Appl Environ Microbiol. 2007;73:1481-1488.
21. Zhao X, Hofte M, Spanoghe P, Rajkovic A, Uyttendaele M. Biofilm-forming Ability of Bacillus thuringiensis Strains from Biopesticides on Polystyrene and their Attachment on Spinach. J Food Protect. 2024;87(8):100321.