Introduction

The growing global demand for sustainable agricultural production has increased interest in replacing or reducing chemical agrochemicals with environmentally friendly alternatives. Among these, plant-associated probiotic bacteria, particularly Pseudomonas and Bacillus species, have gained importance due to their ability to produce antimicrobial metabolites, induce systemic plant resistance, and promote plant growth1,2. Despite their potential, the use of these beneficial microorganisms in the field poses a challenge as their viability and biological activity rapidly decline under unfavorable environmental conditions such as ultraviolet radiation, temperature fluctuations, desiccation, osmotic stress, and soil limitations. This instability not only reduces their effectiveness but also limits the commercial viability of microorganism-based products3.

Encapsulation in biopolymer-based matrices has proven to be a promising strategy to improve the stability and performance of microbial inoculants4. By creating a protective physical barrier, encapsulation shields the cells from environmental stress and allows for controlled, sustained release at the target site4.

The choice of encapsulation method and coating materials is critical to achieving these results. Advanced coating approaches such as layer-by-layer (LbL) and multi-layer encapsulation allow precise control of the capsule architecture and thus the customization of mechanical strength, chemical stability, and release kinetics5. LbL assembly is widely recognized as a versatile and robust platform for the protection and controlled delivery of sensitive bioactive agents, including enzymes, probiotics, drugs, and living cells, extending beyond microbial applications6,7.

Numerous studies have shown that LbL-assembled polysaccharide–protein multilayers significantly enhance structural integrity and functional longevity by regulating interfacial interactions and layer thickness at the nano- to microscale8. Biocompatible and renewable polymers such as sodium alginate, pectin, whey protein, and various native gums have emerged as promising materials for microbial encapsulation9,10,11. Multilayer systems based on natural polysaccharides such as alginate and pectin exhibit improved resistance to mechanical and environmental stresses, as well as more predictable and sustained release profiles12,13,14. Alginate microcapsules have been effectively coated using various strategies with biocompatible polyanions, or neutral polymers, enhancing their protective capacity and functional performance15. Specifically, LbL encapsulation of probiotics with zein nanoparticles and pectin achieved high survival rates (> 95%) and demonstrated that both the number of layers and the outermost composition critically affect viability during heat exposure, storage, and digestion12.

Incorporating proteinaceous components, such as whey protein, further strengthens capsule cohesion through electrostatic interactions, hydrogen bonding, and network formation, resulting in hybrid structures with superior mechanical stability and barrier properties16. Coating airbrush-encapsulated calcium alginate microbeads with regular buttermilk proteins (RBMP) significantly improved the viability, storage stability, and gastrointestinal tolerance of Lactobacillus rhamnosus GG17. Additionally, the use of renewable plant gums as outer or intermediate layers enhances encapsulation efficiency, improves release modulation, and increases sustainability, while providing cost-effective and environmentally friendly formulation options18. Multilayer microcapsules composed of alginate, whey protein, natural gums, and silica/titania nanoparticles effectively encapsulated Bacillus velezensis, preserving its functional activities and providing a biodegradable, cost-effective strategy for sustainable biofertilizers19. Collectively, these findings highlight the strong scientific foundation of LbL-based multilayer encapsulation systems using multiple biopolymers and support their relevance as an effective strategy for bacterial protection and controlled release.

Despite significant advances in microbial encapsulation, the combined use of multiple biopolymers through LbL and multilayer strategies – particularly for the protection and delivery of plant growth-promoting rhizobacteria (PGPR) in plants – remains insufficiently explored, underscoring both the novelty and relevance of this study.

The primary objective of this study was to develop and compare double-layer and multi-layer encapsulation systems based on multiple biopolymers using two distinct coating strategies. Accordingly, alginate–pectin–whey protein–gum formulations were designed and evaluated. This work provides comprehensive insights into the physicochemical properties, release behavior, and potential applications of these systems in the development of next-generation biocontrol products.

The findings of this study have important practical implications for the development of sustainable biofertilizers and next-generation biocontrol products, offering an effective strategy to enhance the viability, stability, and delivery of PGPR under field conditions.

We hypothesize that multilayer encapsulation systems composed of multiple biopolymers will outperform double-layer formulations in enhancing the protection, long-term stability, and controlled release of PGPR, providing a more effective strategy for sustainable biofertilizer and biocontrol applications.

Materials and methods

Materials

Potato dextrose agar (PDA), nutrient agar (NA), and nutrient broth (NB) were used in the preparation of microbial cultures, which were supplied by Merck (Germany). Sodium alginate (Alg) (viscosity = 43 mPa·s, density = 1.6 g·cm⁻³, Mw = 1.39 × 10⁵ g·mol⁻¹, M/G ratio = 1.57) was purchased from Sigma-Aldrich (Germany). Whey protein isolate (WPI, with a protein content of 90%, based on dry matter) was purchased from Alinda (Greece). Pectin (PEC) from citrus peel (galacturonic acid ≥ 74.0%, dried basis) was purchased from Sigma-Aldrich (Germany). Apricot gum (AG) was collected directly from the trunks of Prunus armeniaca trees in Shahrbabak, Kerman, Iran. Hollow mesoporous silica nanoparticle (HMSN) was synthesized in the nanotechnology laboratory of Vali-e-Asr University of Rafsanjan (Rafsanjan, Iran). All chemicals and reagents were of analytical grade and were used without further purification.

Bacterial strains and cultivation procedure

The antagonistic strains Bacillus velezensis VRU1 and Pseudomonas fluorescens T17-4, which originated from the PGPR collection center of the Vali-e-Asr University of Rafsanjan, were stored at − 80 °C in nutrient broth–glycerol (1:2, v/v). Before encapsulation, the cultures were reactivated on NA at 28 °C and then grown in nutrient broth for 24 h, and the resulting suspensions were used to prepare the microcapsules.

Biocontrol and growth-promoting activity assays

Lipase activity

Lipase production was tested on medium containing peptone (10 g/L), NaCl (0.5 g/L), calcium chloride (CaCl₂) (0.1 g/L), and agar (15 g/L) supplemented with sterile Tween 80 (10 ml/L) after sterilization20. The bacterial isolates were spot inoculated and incubated at 28 °C for 72 h. The formation of opaque halos indicated lipase activity.

Cellulase activity

Cellulase activity was determined on carboxymethyl cellulose (CMC) agar medium21. After spot inoculation of 24-hour bacterial cultures, the plates were incubated at 28 °C for five days. The plates were then stained with 1% (w/v) Congo red for 15 min and washed with 1 M NaCl. Clear halos around the colonies indicated cellulase production.

Protease activity

Protease production was determined according to Maurhofer et al.22 using skim milk agar (SMA) with 15 g/L skim milk powder, 0.5 g/L yeast extract, and 12.9 g/L agar. The bacterial isolates were spot inoculated and incubated at 28 °C for 24–48 h. Transparent zones surrounding colonies were considered positive for protease activity.

Hydrogen cyanide (HCN) production

HCN production was assessed according to the method of Alström23 with slight modifications. The bacterial lawn was produced on King’s B agar with 4% (w/v) glycine. A filter paper soaked with 0.5% (w/v) picric acid and 2% (w/v) sodium carbonate solution was placed in the lid of the plate. The plates were sealed with parafilm and incubated at 28 °C for five days. A color change from yellow to orange indicated HCN production.

Phosphate solubilization

Phosphate solubilization was tested with Pikovskaya (PVK) agar medium containing glucose (1% w/v), (NH₄)₂SO₄ (0.05% w/v ), NaCl (0.02% w/v), KCl (0.02% w/v), CaCl₂-2 H₂O (0.01% w/v), MgSO₄-7 H₂O (0.01% w/v), MnSO₄-7 H₂O (0.05% w/v), FeSO₄-7 H₂O (0.05% w/v), yeast extract (0.05% w/v), Ca3(PO4)2 (0.5% w/v) and agar (1.5% w/v), adjusted to pH 7. The bacterial strains were spot-inoculated and incubated for 72 h at 28 °C. Transparent halos around the colonies indicated phosphate solubilization24.

Indole acetic acid (IAA) production

IAA production was quantified according to Patten and Glick25. Bacterial cultures were grown for 72 h in NB broth supplemented with 200 mg/L L-tryptophan and then centrifuged at 1000 rpm for 10 min. Two milliliters of the supernatant were mixed with 4 mL Salkowski reagent (150 mL H₂SO₄, 250 mL distilled water, 7.5 mL 0.5 M FeCl₃-6 H₂O) and kept in the dark for 20 min. The absorbance was measured at 535 nm using a spectrophotometer.

Synthesis of nanoparticles

Hollow mesoporous silica spheres were obtained after hydrolysis and condensation of tetraethoxysilane (TEOS) in a water–ethanol medium (25.5 mL water, 15 mL ethanol) using ammonia (0.5 mL, 25 wt% NH₃ in water) as a catalyst and cetyltrimethylammonium bromide (CTAB, 0.08 g; final concentration 5 mM) as a surfactant26. TEOS (0.5 ml) and CTAB were mixed with the solvent, followed by slow addition of ammonia with stirring (700 rpm) at 25 °C for 3 h. The precipitates were collected by centrifugation, washed with water, and calcined in two steps: 200 °C for 6 h and 600 °C for 6 h.

Characterization of nanoparticles

Transmission electron microscopy (TEM) images were acquired using a Zeiss EM10 microscope. Samples were dispersed in a suitable dispersant and ultrasonicated before a few drops were placed onto Formvar-coated copper grids. Scanning electron microscopy (FESEM) images were acquired using a ZEISS Sigma 300 field emission SEM. Powdered samples were mounted on carbon tape attached to a stub and coated with a thin layer of gold. X-ray diffraction (XRD) patterns were recorded with an X’Pert Pro diffractometer (Panalytical) using Cu Kα radiation (λ = 1.5418 Å); powdered samples were used directly for analysis without prior weighing or drying. Fourier transform infrared (FTIR) spectra were recorded in transmission mode using the KBr pellet method. Samples and KBr were mixed visually, adjusting the ratio according to the sample color to avoid overly dark pellets. No precise weighing, dilution, or drying was performed. FTIR spectra were collected in the range 400–4000 cm⁻¹ with a resolution of 4 cm⁻¹ and 32 scans.

Evaluation of the effect of nanoparticles on the growth of target bacteria

The agar well diffusion method was used to evaluate the inhibitory effect of the nanoparticles used in the formulation. First, a suspension of the bacterial strains was prepared and evenly distributed on nutrient agar plates. At the same time, the suspensions containing the nanoparticles were freshly prepared to minimize experimental errors. Wells with a diameter of 5 mm were then made on the agar surface at a distance of 1 cm from the edge of the petri dish. Subsequently, 40 µL of the nanoparticle suspensions were added to the wells at concentrations of 20 and 40 µg/mL. Sterile distilled water was used as a control. The plates were incubated at 28 °C for 24 h. After incubation, the presence or absence of a clear zone of inhibition around the bacterial colonies was analyzed11.

Encapsulation process

Microcapsule preparation

The microcapsules were produced using two coating strategies: LbL and Multi-layer, using natural polymers such as Alg, AG, WPI, and PEC.

For the LbL method, a sterile 1.5% (w/v) Alg solution and 4% (w/v) WPI were prepared as a core matrix supplemented with 40 µg/mL HMSNs. The bacterial suspension (2 × 1010 CFU/mL) was added to this solution and mixed with a magnetic stirrer for 30 min. The polymers–bacteria mixture was then extruded with a syringe into a 2% (w/v) CaCl₂ solution, where the resulting beads were stirred for 30 min to promote ionic cross-linking. The hardened beads were then immersed in an AG solution as a second coating layer and gently shaken at 100 rpm for 30 min at room temperature. To strengthen the coating, the beads were immersed again in 2% CaCl₂ for 30 min. The finished microcapsules were rinsed with sterile physiological saline solution (0.9% NaCl) and dried at 40 °C in the laboratory oven.

In the multilayer method, the core formulation consisted of 1.5% (w/v) Alg, 4% (w/v) WPI, 40 µg/mL HMSNs, and 2 × 1010 CFU/mL bacterial suspension. This mixture was extruded in 2% CaCl₂, and the beads were stirred for 30 min to gel. The beads were then transferred to an AG solution (second coating layer) and stirred for 30 min, followed by immersion in 2% CaCl₂ for 30 min. The beads were then coated with 2% (w/v) PEC (third coating layer) with gentle stirring for 30 min, and then crosslinked again in 2% CaCl₂ for 30 min. The resulting microcapsules were washed with 0.9% NaCl and dried at 40 °C.

The encapsulation strategy is based on well-established alginate–calcium gelation and LbL coating methods for microbial encapsulation. The concentrations of Alg (1.5% w/v), CaCl₂ (2% w/v), and WPI (4% w/v) were selected to ensure proper bead formation, mechanical stability, and bacterial viability. The concentrations of AG and PEC were chosen for their film-forming ability and compatibility with the alginate matrix. These conditions were optimized through preliminary experiments to obtain stable microcapsules with uniform size and high bacterial survival.

After preparation, the morphology and structural properties of the microcapsules were characterized using SEM, FTIR, and XRD. SEM observations were performed with a ZEISS Sigma 300 field emission scanning electron microscope at different magnifications to evaluate capsule shape, surface morphology, and layer integrity. FTIR spectra and XRD patterns of the microcapsules were recorded under the same conditions as those used for nanoparticle characterization.

Encapsulation efficiency of microcapsules

The microcapsules (1 g) were suspended in 9 mL of phosphate buffer (0.2 M, pH 7.4) and incubated for 1 h at 28 °C in a shaker to release the entrapped cells27. Cell viability was determined using the standard plate count method. Appropriate 10-fold serial dilutions were prepared and plated on nutrient agar plates. Colony-forming units (CFU) were counted after 24 h of incubation at 28 °C. The encapsulation efficiency (EE) was calculated according to Eq. (1):

$$\:EE\:\left(\%\right)=\:\frac{N}{{N}_{0}}\:\times\:100$$
(1)

Where N is the number of viable entrapped cells released from 1 g microcapsules, and N0 is the number of free cells added to the biopolymer mix.

Moisture content of microcapsules

The moisture content was determined by weighing 1 g of the wet microcapsules, drying at 40 °C and weighing again. It was calculated using Eq. (2)28:

$$\:\text{M}\text{o}\text{i}\text{s}\text{t}\text{u}\text{r}\text{e}\:\text{c}\text{o}\text{n}\text{t}\text{e}\text{n}\text{t}\:\left(\text{\%}\right)=\:\frac{\text{W}\text{b}-\text{W}\text{a}}{\text{W}\text{b}}\:\times\:100$$
(2)

Wb and Wa are the weights (g) of microcapsules before and after drying, respectively.

Swelling ratio of microcapsules

The swelling ratio was determined by immersing 1 g of the dried microcapsules in phosphate buffer (0.2 M, pH 7.4) for 24 h, blotting off excess liquid, and weighing the swollen beads as shown in Eq. (3)29:

$$\:\text{S}\text{w}\text{e}\text{l}\text{l}\text{i}\text{n}\text{g}\:\text{r}\text{a}\text{t}\text{i}\text{o}\:\left(\text{\%}\right)=\:\frac{\text{W}\text{b}-\text{W}\text{a}}{\text{W}\text{b}}\:\times\:100$$
(3)

Wb and Wa are the weights (g) of swelling and dried microcapsules, respectively.

Release behavior of bacterial strains

To evaluate the release of the bacteria, 1 g of microcapsules was placed inside a 12 kDa dialysis bag, with both ends partially closed using clips to contain the capsules while allowing gradual exchange with the surrounding phosphate buffer. The dialysis bag was immersed in 400 mL of phosphate buffer (0.2 M, pH 7.4) at 28 ± 2 °C. At intervals of 5 days up to 60 days, aliquots of the buffer were sampled to monitor the release of viable bacteria. Upon immersion, the microcapsules swelled, and bacteria were gradually released through the openings at the bag closures into the buffer, enabling assessment of release kinetics.

Long-term stability test

The microencapsulated bacteria were stored for up to 6 months in sterile microtubes, at room temperature (25 ± 0.5 °C). The viability of the bacterial strains was determined using the plate count method. One gram of microcapsules (with an initial concentration of 2 × 1010 CFU/mL) was suspended in 9 mL of sterile phosphate buffer (0.2 M, pH 7.4) and allowed to disintegrate for 1 h. The number of viable cells was determined by serial dilution and plating on nutrient agar, followed by incubation at 28 °C for 48 h. Results were expressed as mean ± SD of three independent experiments.

Statistical analysis

All experiments were performed in triplicate (n = 3), and results are expressed as mean ± standard deviation (SD). Statistical analyses were conducted using one-way analysis of variance (ANOVA), followed by Tukey’s multiple comparison post hoc test, with SAS software version 9.1 (SAS Institute, Inc., Cary, NC, USA). Differences among treatments were considered statistically significant at p < 0.05 and are denoted as follows: ns, p > 0.05; *, p < 0.05; **, p < 0.01.

Results and discussion

Plant growth-promoting and biocontrol characteristics

Both P. fluorescens T17-4 and B. velezensis VRU1 exhibited multiple plant growth-promoting and biocontrol traits (Fig. 1). Both strains produced hydrolytic enzymes, including cellulase, protease, and lipase, solubilized inorganic phosphate, synthesized siderophores, released HCN; and generated IAA.

Fig. 1
figure 1

Comparative analysis of plant growth–promoting and enzymatic traits of Pseudomonas fluorescens T17-4 (upper row) and Bacillus velezensis VRU1 (lower row), including, (a) Production of cellulase, (b) Production of protease, (c) Production of lipase, (d) Phosphate solubilization, (e) Production of siderophore, (f) Production of HCN, (g) Production of IAA.

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Encapsulation process

Characterization of nanoparticles

The synthesized HMSN exhibited a well-defined spherical morphology, uniform size distribution and smooth surface texture. SEM images (Fig. 2a) confirmed that the particles maintained their structural integrity even after high-temperature calcination and showed no significant collapse or deformation. The particle diameters ranged from 400 to 480 nm with a narrow distribution, reflecting the successful control of the synthesis process. Such uniformity is expected to play a crucial role in improving the surface properties and functional performance of these materials in various applications. The EDS analysis (Fig. 2c) confirmed silicon (54.7 wt%) and oxygen (45.3 wt%) as the dominant elements, which is consistent with the SiO₂ stoichiometry. Minor gold peaks were attributed to the conductive coating applied during SEM imaging and did not correspond to the actual composition of the nanoparticles. Moreover, elemental mapping (Fig. 2b) showed the homogeneous spatial distribution of Si and O, confirming the chemical purity and structural uniformity of the synthesized MSNs. TEM images (Fig. 2d) showed worm-like mesoporous channels distributed across the particle shells, and parallel lattice fringes, confirming the structural order at close range. Statistical analysis of the TEM images revealed an average particle diameter of ~ 350 nm with a narrow size distribution, which is consistent with the SEM results. The FTIR spectrum (Fig. 2e) further confirmed the structural framework and surface functionalities. A broad band at 3500–3400 cm⁻¹ corresponded to the stretching vibration of hydroxyl groups (Si–OH) and adsorbed water molecules, while the band at 1628 cm⁻¹ was assigned to the bending vibration of physisorbed water. A strong peak at 1086 cm⁻¹, attributed to asymmetric Si–O–Si stretching, and weaker bands at 800 and 462 cm⁻¹ corresponding to symmetric and bending vibrations, confirm the formation of a three-dimensional siloxane network30. The XRD pattern (Fig. 2f) shows a pronounced low-angle reflection, which is characteristic of mesoporous structures, and indicates a short-range ordering of the pore system. The relatively low peak intensity was attributed to the hollow interiors of the particles and the limited long-range ordering of the mesostructured26.

Fig. 2
figure 2

(a) SEM image, (b) SEM-EDS mapping pattern, (c) EDS analysis, (d) TEM image, (e) FTIR spectrum, and (f) XRD pattern of the hollow mesoporous silica nanoparticles.

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The effect of nanoparticles on bacterial strains growth

The effect of HMSN on two bacterial strains, VRU1 and T17-4, was investigated. The observations showed that neither concentration inhibited bacterial growth, as no zones of inhibition were detected (Fig. 3). These results indicate a lack of antibacterial activity of the nanoparticles against these strains.

Fig. 3
figure 3

Antibacterial activity of hollow mesoporous silica nanoparticles (HMSNs) against bacterial strains VRU1 and T17-4 evaluated by the agar well diffusion method. HMSNs were tested at concentrations of 20 and 40 µg/mL. Control wells contained no nanoparticles (0 µg/mL).

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Morphological characterization of microcapsules

Figure 4 presents the morphological characteristics of microcapsules fabricated with different wall architectures. The optical microscopy image of the double-layer system (Fig. 4A) shows well-defined capsules with a clear core–shell boundary. However, SEM images (Fig. 4B, C) show a rough and heterogeneous surface morphology composed of relatively small, loosely packed particles. These surface features suggest a porous shell structure, which may facilitate the diffusion of encapsulated compounds while potentially reducing the mechanical integrity of the capsule wall.

In contrast, multilayer microcapsules display a markedly different morphology. The optical image (Fig. 4E) indicates a more compact shell compared to the double-layer system, while SEM observations (Fig. 4F, G) show smoother, denser, and more continuous surfaces. Although shell thickness was not measured directly, the observed surface compactness suggests the formation of a more cohesive shell structure. This behavior can be attributed to enhanced intermolecular interactions between successive layers, which promote the development of a dense and interconnected network, as commonly reported for multilayer encapsulation systems31,32.

Microcapsule size distributions were estimated from measurements of 13 individual capsules obtained from SEM images. Although the sample size is limited, the results are consistent with the qualitative morphological observations. Double-layer capsules have smaller and more uniform diameters, while multilayer capsules display a broader and more heterogeneous size distribution. These trends are shown in the corresponding size distribution plots (Fig. 4), where double-layer capsules (Fig. 4D) display a narrower diameter range than multilayer capsules (Fig. 4H). It should be noted that these size distributions are approximate and do not constitute statistically robust analyses.

The observed structural differences between double- and multilayer systems are expected to influence their functional performance. The more porous shell morphology of double-layer capsules may contribute to lower encapsulation efficiency and faster release in aqueous environments, whereas the denser and more cohesive shell structure of multilayer capsules is likely associated with improved retention and a more controlled release profile, consistent with previous reports on multilayer encapsulation architectures.

Fig. 4
figure 4

Optical microscopy images and SEM micrographs of double-layer microcapsules (upper row, A–D) and multi-layer microcapsules (lower row, E–H), illustrating overall morphology and surface characteristics. Histograms showing the estimated size distribution of double-layer and multi-layer microcapsules based on measurements of n = 13 individual capsules from SEM images.

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X-ray diffraction analysis

The XRD patterns of Alg, WPI, PEC, AG, and the corresponding assembled microcapsules are shown in Fig. 5. Sodium alginate displayed a broad diffraction halo without distinct sharp peaks, indicating a predominantly amorphous structure. The absence of well-defined crystalline reflections suggests a lack of long-range molecular order, attributed to the flexible polymer backbone, irregular chain packing, and random orientation of alginate macromolecules, as commonly reported for alginate-based materials. Similarly, AG displayed a diffuse and poorly resolved diffraction profile, confirming its amorphous nature. This behavior is typical of natural plant gums composed of heterogeneous polysaccharide chains with limited structural regularity33,34,35. The XRD pattern of WPI was also dominated by a broad halo, reflecting its largely amorphous structure caused by the disordered arrangement of protein chains and the absence of periodic crystalline domains36,37. In contrast, PEC exhibited several sharp and intense diffraction peaks at specific 2θ values, indicating a semi-crystalline structure. These crystalline features result from localized chain ordering and intermolecular associations within pectin domains, as previously reported in the literature38,39.

After layer-by-layer assembly, both Alg/WPI–AG and Alg/WPI–AG–PEC microcapsules showed a marked decrease in XRD peak intensity and sharpness, along with the appearance of broad amorphous halos, indicating a significant loss of crystalline order. Similar reductions in crystallinity have been reported for Alg–WPI systems and alginate–pectin complexes, where strong electrostatic interactions and hydrogen bonding disrupt the ordered crystalline domains of the individual biopolymers during complexation40,41. In particular, the weakening or disappearance of characteristic crystalline reflections of PEC has been attributed to conformational rearrangements and polymer chain entanglement induced by biopolymer interactions, resulting in a more homogeneous and predominantly amorphous matrix42. This increase in amorphous character is advantageous for encapsulation applications, as amorphous biopolymer matrices generally provide greater flexibility, improved molecular dispersion, higher encapsulation efficiency, and more controlled release of bioactive compounds compared to semi-crystalline systems, a phenomenon commonly observed in polyelectrolyte multilayer assemblies41.

Fig. 5
figure 5

X-ray diffraction (XRD) patterns of alginate (Alg), whey protein isolate (WPI), Pectin (PEC), apricot gum (AG), alginate/whey protein-apricot gum microcapsule (Alg/WPI-AG), and alginate/whey protein-apricot gum-pectin microcapsule (Alg/WPC-AG-PEC).

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FTIR analysis

The FTIR spectra of Alg, AG, WPI, PEC, and their microcapsules are illustrated in Fig. 6. The FTIR spectrum of AG clearly shows characteristic polysaccharide features. A broad and intense absorption band at 3425 cm⁻¹ was assigned to O–H stretching vibration .In addition, the absorption in the range 2932 cm⁻¹ was assigned to the –CH₂ stretching vibrations. The distinct band at 1609 cm⁻¹ was attributed to carboxylate groups, while the strong absorption at ~ 1033 cm⁻¹ corresponded to C–O stretching and glycosidic bonds (e.g., arabinosyl residues). These observations are consistent with the reports describing plant gums with hydroxyl, methylene, and carbonyl functions and confirm that apricot gum has structural features similar to other galactomannans43,44,45. The FTIR spectrum of WPI shows distinct bands associated with protein structures. The broad absorption near 3292 cm⁻¹ was attributed to O–H stretching vibrations associated with amide groups. The peaks in the 2920–2970 cm⁻¹ region were assigned to methylene (–CH₂) stretching. The characteristic absorption at 1649 cm⁻¹ corresponded to the amide I band (C = O stretching), while the band at 1532 cm⁻¹ was attributed to the amide II band (N–H bending). These results are in excellent agreement with previously reported FTIR data for whey protein46. The spectrum of Alg showed characteristic peaks corresponding to its functional groups. The broad band at 3446 cm⁻¹ was assigned to the O–H stretching vibrations. The absorptions at 1613 and 1429 cm⁻¹ were assigned to the asymmetric and symmetric stretching of carboxylate groups (COO⁻), respectively. A strong peak at ~ 1021 cm⁻¹ corresponded to C–O stretching vibrations in the polysaccharide backbone47. The FTIR spectrum of PEC showed the characteristic features of polysaccharides. A broad band in the region of 3300–3400 cm⁻¹ was attributed to O–H stretching. The band at 2923 cm⁻¹ was assigned to the –CH stretching vibrations. A prominent band at 1601 cm⁻¹ corresponded to carboxylate groups, while the peak around 1014 cm⁻¹ was related to C–O–C stretching in glycosidic bonds48. Finally, the FTIR spectra of the double-layer and multi-layer microcapsules displayed combined features of their individual components, indicating successful assembly of the multi-layer structures. In both systems, the presence of broad O–H stretching bands, carboxylate-related absorptions, and glycosidic vibrations confirmed polysaccharide–polysaccharide interactions, while the amide I and amide II bands indicated the incorporation of WPI into the microcapsule matrix. FTIR spectroscopy does not directly provide spatial information about the exact positioning of individual layers49. The identification of pectin as the outermost layer in the multi-layer microcapsules is based on the layer-by-layer fabrication process. However, the FTIR spectrum of the multi-layer microcapsules shows increased intensity and slight shifts in the carboxylate band (≈ 1617 cm⁻¹) and the C–O–C stretching region (≈ 1011 cm⁻¹), which are characteristic of pectin. These changes, compared to the double-layer system, indicate the successful incorporation of pectin and its interaction with the underlying layers. Additionally, increased band broadening and intensity in the O–H and carboxylate regions suggest the formation of hydrogen bonds and electrostatic interactions among WPI, Alg, AG, and PEC. These intermolecular interactions contribute to the formation of a denser and more stable matrix49. The addition of a PEC layer in the multi-layer microcapsules is therefore expected to enhance structural integrity and improve the controlled release performance of encapsulated bioactive compounds50,51.

Fig. 6
figure 6

FT-IR spectra of the multi-layer and double-layer microcapsule, whey protein isolate (WPI), pectin (PEC), apricot gum (AG), and alginate (Alg).

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Efficiency of encapsulation

Encapsulation efficiency was significantly affected by the coating formulation and layering strategy (p < 0.05) (Table 1). The double-layer system (Alg + WPI-AG) achieved efficiencies of 83% for VRU1 and 80% for T17-4, with no significant difference between strains. Although these values indicate relatively high entrapment, they suggest that the double-layer coating does not provide sufficient barrier properties to fully prevent bacterial leakage. In contrast, the multi-layer system (Alg + WPI-AG-PEC) significantly improved encapsulation efficiency, reaching 90.66% for VRU1 and 90% for T17-4. The incorporation of pectin likely enhanced capsule compactness through additional hydrogen bonding and electrostatic interactions, strengthening the hydrogel network and reducing porosity.

Swelling percentage and moisture content

The results showed that moisture content was significantly affected by wall composition (p < 0.05) (Table 1). The double-layered capsules made with Alg + WPI–AG had the lowest moisture values (40.50–40.73%), while adding PEC in the three-layered system (Alg + WPI–AG–PEC) significantly increased moisture content to 63.80–64.69%. The strong hydrophilic nature and water-retention capacity of PEC likely contributed to this increase by promoting the formation of a more porous hydrogel network that can bind higher amounts of water.

In contrast, swelling behavior showed no significant differences among treatments (p > 0.05) (Table 1), with values ranging narrowly from 119.67% to 121.12%. This suggests that although the presence of PEC modified the water content of the microcapsules, it did not substantially alter the overall swelling dynamics, which is consistent with previous findings showing that increased pectin concentration enhances gel strength but does not necessarily increase swelling capacity.

Long-term stability

The survival of encapsulated cells over six months of storage showed that wall composition was crucial for long-term stability. The double-layered formulations (Alg + WPI–AG) maintained viability at 80.17% (VRU1) and 79.92% (T17-4), while the addition of pectin in the multi-layered system (Alg + WPI–AG–PEC) significantly improved survival to 90.27% (VRU1) and 91.60% (T17-4) (p < 0.05) (Table 1). The enhanced stability of the multi-layered capsules can be attributed to the hydrophilic and film-forming properties of PEC, which may improve water retention and reduce oxidative or desiccation stress during storage. In addition, PEC is known to form a denser polymeric network through calcium crosslinking, thereby decreasing oxygen permeability and protecting bacterial cells from harmful environmental factors.

Table 1 Moisture, swelling, encapsulation efficiency, and 6-month stability of B. velezensis VRU1 and P. fluorescens T17-4 in double- and multi-layer microcapsules.
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Release behavior

The release behavior of encapsulated bacteria was strongly influenced by capsule architecture (Fig. 7). Although the release profiles are shown as continuous curves, statistical analysis (one-way ANOVA followed by Tukey’s post hoc test, P < 0.05) confirmed significant differences between double-layered and multi-layered capsules during the initial stage (0–20 days). As shown in Fig. 7, multi-layered capsules exhibited significantly lower bacterial release compared to double-layered capsules during this period. This reduced release can be attributed to the denser pectin layer in multi-layered capsules, which decreases pore size and limits diffusion of the encapsulated bacteria. This reduction in burst release has been widely reported in multi-layered hydrogel systems and is considered beneficial for early protection of probiotics during storage and processing. During the mid-term phase (20–40 days), both capsule types reached maximum counts of approximately 9–10 log10 CFU. g⁻¹, indicating that diffusion through hydrated matrices governs release at this stage. In the late phase (40–60 days), clear differences emerged. Double-layer capsules showed a decline in CFU counts, likely due to structural degradation and reduced bacterial survival, while multi-layer systems maintained higher release and even slight increases at certain points. This prolonged stability highlights the protective effect of the additional PEC barrier, consistent with previous findings that multi-layer encapsulation enhances long-term survival by reducing matrix erosion and shielding bacteria from environmental stress. No significant differences were observed between bacterial strains, suggesting that capsule design, rather than bacterial strain, primarily determines release dynamics under the tested conditions. Collectively, these results confirm that while double-layer capsules provide faster initial release, multi-layer systems ensure prolonged protection and sustained delivery, making them more suitable for various applications. In functional foods and nutraceuticals, multi-layer capsules help maintain bacterial viability during storage and processing by reducing early burst release. In pharmaceutical or feed applications, these systems provide controlled delivery over extended periods, enhancing efficacy under harsh environmental conditions. The additional pectin layer acts as a barrier against diffusion and environmental stresses, making multi-layer capsules more advantageous than double-layer capsules when long-term stability and gradual release are required.

Fig. 7
figure 7

Release of P. fluorescens T17-4 and B. velezensis VRU1 from double-layer and multi-layer microcapsules in phosphate buffer over 60 days. Data are presented as mean ± SD (n = 3). Statistical differences between treatments at each time point were determined by one-way ANOVA followed by Tukey’s post hoc test (P < 0.05).

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Discussion

The findings of this study highlight the mechanistic advantages of using P. fluorescens T17-4 and B. velezensis VRU1 as multifunctional bioinoculants. The synergistic activities of hydrolytic enzymes, phosphate solubilization, siderophore production, HCN release, and IAA synthesis not only promote nutrient acquisition and root development but also actively modulate the rhizosphere microbial community, thereby suppressing phytopathogens52,53,54,55,56. These mechanisms suggest that, while the bacterial strains play a crucial role in promoting plant growth and controlling diseases, the encapsulation matrix provides protection and sustained release, both essential for their effectiveness in field applications.

Despite their intrinsic capabilities, free bacterial inoculants often experience rapid population decline due to environmental stresses such as osmotic fluctuations, oxidative stress, temperature variability, and competition with indigenous microbiota. Encapsulation creates a physical and chemical microenvironment that mitigates these stresses, enhances bacterial resilience, and prolongs metabolic activity. In particular, multi-layer capsules with pectin as the outermost layer improve structural cohesion and reduce matrix porosity, thereby limiting the diffusion of extracellular enzymes and metabolic by-products that could otherwise compromise bacterial viability19,57,58,59.

Structural analyses (SEM, XRD, FTIR) revealed that interpolymer hydrogen bonding, electrostatic interactions, and molecular entanglement among alginate, whey protein, apricot gum, and pectin form a dense, amorphous network, which enhances mechanical robustness and provides a semi-permeable barrier. This barrier selectively modulates water and nutrient diffusion, protects against desiccation, and buffers oxidative damage, thereby maintaining bacterial metabolic activity during extended storage33,50,51. The addition of mesoporous silica nanoparticles further reinforces capsule rigidity, serves as a scaffold for structural integrity, and increases the water retention capacity of the hydrogel matrix, collectively creating a biocompatible microenvironment that mimics soil pore networks60.

Functionally, multi-layered capsules modulate the release kinetics of encapsulated bacteria through steric hindrance and matrix density. The denser pectin outer layer delays burst release during early storage, preventing rapid depletion of bacterial populations and maintaining bioactivity during the critical initial phase of field application61,62,63,64. During prolonged storage, controlled diffusion through the hydrated matrix ensures sustained release, which is crucial for maintaining effective rhizosphere colonization and continuous expression of PGP and biocontrol functions. This mechanistic insight explains the superior long-term viability (> 90% after six months) observed for multi-layer formulations compared to double-layer systems.

Collectively, these results demonstrate that the encapsulation matrix, rather than the bacterial strain, is the primary determinant of functional efficacy. By integrating hierarchical polymer layers with nanoparticles, the system provides mechanical and chemical protection, stabilizes bacterial metabolism, regulates release, and optimizes interactions with plant roots. This strategy offers a scalable platform for developing next-generation biofertilizers to address field-level challenges in sustainable agriculture. Field-level limitations for microbial inoculants, such as temperature fluctuations, desiccation, UV exposure, osmotic stress, competition with native microbiota, and uneven rhizosphere colonization, are effectively mitigated by the multi-layer encapsulation approach, ensuring gradual and sustained bacterial delivery over extended periods. This reduces the need for repeated inoculations, promotes consistent expression of beneficial traits, and enhances rhizosphere colonization.

Limitations and future perspectives

This study aimed to evaluate the viability, stability, and release behavior of PGPR encapsulated within double- and multi-layer biopolymer microcapsules. Although the results demonstrate the effectiveness of multilayer capsule architectures in enhancing bacterial protection and sustained release, several limitations should be acknowledged.

First, while SEM, FTIR, and XRD analyses were used to characterize the nanoparticles and microcapsules, the spatial distribution of HMSN within the microcapsule core was not quantitatively assessed. As the nanoparticles were incorporated exclusively into the core matrix, advanced imaging techniques such as elemental mapping or three-dimensional imaging would be required for detailed localization, which is recommended for future studies. Second, bacterial release behavior was interpreted based on experimentally observed and statistically supported trends, although quantitative mechanistic modeling was not applied. Future incorporation of mathematical modeling could provide a more comprehensive understanding of release mechanisms. Third, despite the suitable qualitative flow behavior and mechanical integrity of the polysaccharide–protein core formulation, detailed rheological characterization was beyond the scope of this work. Such analysis would be beneficial for formulation optimization and scale-up considerations. Finally, the experiments were conducted under controlled laboratory conditions. Future studies involving soil-based, greenhouse, and field evaluations will be essential to validate the performance of the encapsulated PGPR under practical agricultural environments.

Overall, despite these limitations, the present study provides a robust foundation for the development of multilayer biopolymer microcapsules as effective delivery systems for beneficial rhizobacteria and outlines clear directions for future research.