Tenoxicam-loaded bioglass/chitosan composites for bone tissue engineering: in vitro characterization, sustained drug release, and antimicrobial activity

Abstract

The development of multifunctional biomaterials that combine structural support with therapeutic functions is a key challenge in bone tissue engineering. Bioglass and chitosan are widely studied for their bioactivity and biocompatibility, while non-steroidal anti-inflammatory drugs (NSAIDs) such as Tenoxicam provide pain relief and inflammation control. We synthesized novel Tenoxicam-loaded Bioglass/Chitosan (TXC/BG/Cs) composites via the sol–gel process, incorporating 1, 2, and 3 wt% drug concentrations. The composites were characterized by FTIR, XRD, SEM, and EDX before and after immersion in simulated body fluid (SBF) to evaluate physicochemical properties and bioactivity. Sustained drug release was monitored for 33 days using UV–Vis spectroscopy. Antibacterial activity was assessed against Gram-positive (Staphylococcus aureus, S. haemolyticus) and Gram-negative (Escherichia coli, Klebsiella pneumoniae) bacteria using the agar well diffusion method. FTIR and XRD confirmed the formation of hydroxyapatite layers on composite surfaces after immersion in SBF, indicating strong bioactivity. SEM revealed a porous microstructure favorable for osteointegration, while EDX confirmed increased Ca and P deposition. The composites exhibited sustained Tenoxicam release following zero-order kinetics, with C3 (3 wt%) showing the highest cumulative release. Antimicrobial testing demonstrated significant inhibition zones against both Gram-positive and Gram-negative strains. These findings suggest that TXC/BG/Cs composites integrate drug delivery, antibacterial functionality, and bone-bonding capability, making them promising candidates for bone tissue engineering and post-surgical implants.

Introduction

Bone defects caused by trauma, tumors, infection, or degenerative diseases present major clinical challenges, especially when inflammation and bacterial contamination are present. Conventional systemic therapies often fail to achieve sufficient local drug concentrations, resulting in delayed healing or implant failure. Complex injuries require treatments that balance mechanical stabilization, enhanced regeneration, and infection control. Despite progress in biomaterials, integrating mechanical support, bio-stimulation, and inflammation management remains a significant challenge in bone tissue engineering. Non-steroidal anti-inflammatory drugs (NSAIDs) are widely used to alleviate pain and inflammation by inhibiting prostaglandin synthesis. Incorporating these agents into multifunctional biomaterials offers a promising approach to improve local healing, osteogenesis, and infection prevention, underscoring the need for advanced bioactive materials at the interface of biophysics and regenerative medicine^1–3. Bioglass is renowned for its exceptional bioactivity, characterized by rapid hydroxyapatite (HA) layer formation that enables strong chemical bonding with bone tissue. Composed primarily of silica, sodium, calcium, and phosphorus, it exhibits excellent biocompatibility, osteo-conduction, and mineralization promotion, alongside angiogenic and antimicrobial properties, making it a promising biomaterial for bone repair and regenerative therapies despite its mechanical brittleness in hydrated environments^4–6.

Chitosan, a natural polysaccharide derived from chitin, has attracted considerable attention as a complementary material due to its biocompatibility, biodegradability, antimicrobial properties, and excellent film-forming capability. When combined with bioglass, chitosan can improve mechanical integrity, enhance the handling characteristics of the composite, and support controlled drug release. Several studies have explored bioglass/chitosan systems for tissue engineering and drug delivery, demonstrating improved structural stability and enhanced bioactivity⁷.

NSAIDs such as Tenoxicam are widely used to control pain and inflammation associated with bone injuries and orthopedic procedures. However, systemic administration is often limited by short half-life, gastrointestinal side effects, and poor localization at the injury site. Local delivery of Tenoxicam through a bioactive composite matrix offers several advantages: achieving sustained therapeutic concentrations, minimizing systemic toxicity, and supporting an optimal healing environment. Previous investigations into NSAID-loaded biomaterials have shown promising results in reducing inflammation and promoting regeneration, yet reports on Tenoxicam-loaded bioglass/chitosan composites remain scarce, representing a significant gap in the current literature^8–10.

This study aims to develop a novel multifunctional composite by combining chitosan polymer with bioglass and loading it with tenoxicam, resulting in a topical therapeutic system designed to accelerate bone regeneration while effectively reducing inflammation. The composite (TXC/BG/Cs) was synthesized via the sol–gel process to enable sustained and controlled drug release at the injury site. Comprehensive characterization was performed using XRD, FTIR, SEM, and EDX to evaluate the crystal structure, functional groups, morphology, and elemental distribution before and after immersion in simulated body fluid (SBF). Bioactivity was assessed by monitoring hydroxyapatite formation, and the antibacterial activity of the drug-loaded composites was tested against clinically relevant Gram-positive (Staphylococcus aureus, Staphylococcus hemolyticus) and Gram-negative (Escherichia coli, Klebsiella pneumoniae) bacteria using the agar diffusion method. Drug release kinetics were analyzed by UV spectroscopy. By integrating bioactivity, anti-inflammatory drug delivery, and antimicrobial properties, this work presents a promising composite material tailored to meet key clinical demands in bone tissue engineering and post-surgical implant applications.

Materials and methods

Calcium nitrate tetrahydrate (Ca(NO₃)₂·4H₂O, Mw = 236.15 g/mol), Tetraethyl orthosilicate (TEOS, C₈H₂₀O₄Si, Mw = 208.33 g/mol), Tetraethyl phosphate (TEP, C₆H₁₅O₄P, Mw = 182.15 g/mol), and 2 M nitric acid (HNO₃) were provided by Merck Inc. (Darmstadt, Germany). Mw = 1,526,454 g/mol of chitosan nanoparticles (Cs), [C₆H₁₁NO₄]n, was purchased from Across Organic Ltd. (New Jersey, USA). The source of the 20 mg Tenoxicam supplied as Epicotil^® lyophilized vial (C₁₃H₁₁N₃O₄S₂, Mw = 337.4 g/mol) and was purchased from (E.I.P.I. Co., Cairo, Egypt) was used as the active pharmaceutical ingredient. Sigma-Aldrich provided additional chemicals for the preparation of simulated bodily fluid (SBF) (St. Louis, MO, USA).

Synthesis of bioactive glass (BG)

Following the procedure outlined by^11–17, a bioactive glass was created using the sol-gel technique and had a chemistry consisting of 50 weight% SiO₂, 35 weight% CaO, and 15 weight% P₂O₅. In detail, 18.96 ml of TEOS, and 6.94 ml of 2 M Nitric acid (HNO₃) were combined; to help with the acid hydrolysis of TEOS and the combination was left to react for 60 min. As illustrated in Fig. 1 and Table 1, the subsequent reagents were then introduced one after the other and allowed to react for one hour each: add 3.31 ml of TEP and 15.04 g of the calcium nitrate tetrahydrate.

Fig. 1.

Flowchart for Bioglass/chitosan composite preparation steps.

Table 1.

Chemical composition in 1 g of samples prepared by sol–gel.

Sample	TEOS	TEP	(Ca (NO3)2.4H2O)	H2O	HNO3	Chitosan	Tenoxicam
BG	1.896 ml	0.33 ml	1.50398 g	1.8 ml	0.694 ml	0	0
C1						0.1 g	10 mg
C2						0.1 g	20 mg
C3						0.1 g	30 mg

To finish the hydrolysis, the mixture was constantly agitated for one and a half hours. The water evaporated by heating it to 120 °C for three days. The glass (BG) powder was then obtained by heating the powder for three hours at 800 °C to eliminate harmful nitrate ions and add calcium to the silicate network. Following deagglomeration with a mortar and pestle, the BG powders were sieved to a 120 μm particle size¹⁸.

Preparation of tenoxicam-loaded BG/Cs composite

A flowchart showing the steps involved in creating the BG/Cs composite loaded with Tenoxicam is shown in Fig. 2. To make a Cs solution, 0.3 g of Cs polymer powder was dissolved in 30 ml of distilled water with 0.3 ml of acetic acid to distribute into three samples, and the mixture was agitated for three hours.

Fig. 2.

Flowchart for preparation of the BG/Cs composite with Tenoxicam loaded and investigating steps.

In order to create the BG/Cs composite, which has a molar ratio of 25:1 weight%, 2.5 g of the produced BG powder was combined with 40 ml of distilled water for each sample and then added to the Cs solution. Subsequently, the BG/Cs mixture was blended with Tenoxicam at different weight percentages of 1%, 2%, and 3%. For three days, the resultant composite was dried at 120 °C.

To create discs of each sample, the dry powders (0.5 g each) were crushed for 5 min and compressed at 750 psi. The corresponding designations for the composites were C1 (1TXC/BG/Cs), C2 (2TXC/BG/Cs), and C3 (3TXC/BG/Cs), respectively¹⁹. FTIR, XRD, SEM, and other methods were used to analyze the materials, and UV-VIS spectroscopy was used to investigate the release profile of Tenoxicam.

Bioactivity study

The bioactive characteristics of the TXC-loaded BG/Cs composite were investigated in vitro by immersion of the samples in simulated body fluid (SBF), following Kokubo’s standard methodology². Each 0.75 g sample was immersed in 50 ml of the SBF solution at 37 °C for thirty-three days in a water bath with constant stirring to simulate the movement of blood within the tissues of living organisms. After being removed from the SBF, the samples were cleaned with distilled water and allowed to air dry.

Characterization

Before and after being submerged in SBF, the powder samples of BG and TXC-loaded BG/Cs composites were examined using a variety of physicochemical methods. Using a Bruker D2 Phaser Benchtop XRD Diffractometer from Germany and a copper target (Cu Kα = 1.54060 Å) and a nickel filter, the X-ray powder diffraction (XRD) investigation determined the phases and whether the TXC-loaded BG/Cs composite surfaces were crystalline or amorphous. The XRD measurements used a step size of 0.014° in 1 s and spanned a 2θ range from 5° to 80°. Using a Nicolet iS10 (Thermo Scientific, USA), Fourier Transform Infrared Spectroscopy (FTIR) was utilized to identify the functional groups in the BG/Cs composite loaded with Tenoxicam drug. The samples were finely powdered, mixed with KBr at a 1:100 ratio, and pressed into discs under a pressure of 10 tons/cm². With a resolution of 4 cm⁻¹, FTIR spectra were captured in absorbance mode between 400 and 4000 cm⁻¹. The samples were crushed, combined, and formed into pellets. Each sample included 200 mg of KBr and 1 mg of material. FEG-SEM: scanning electron microscopy using a Field Emission Gun (XL30, Philips) was used to analyze the surface morphology at a 30 kV accelerating voltage. After being mounted on carbon-stickered stubs, the specimens were examined under a microscope. Energy dispersive X-ray spectroscopy (EDX Analysis) was connected to the SEM (30 mm² Si (Li) R-RSUTW detector) at 15 kV for chemical analysis. The calcium (Ca) and phosphorus (P) intensities in the composites were compared using EDXA before and after they were submerged in SBF. A series of TXC drug reference solutions with progressively lower concentrations was used to calibrate the UV-VIS absorption spectroscopy equipment (JASCO v-630) at the maximum wavelength (λ) of 370 nm, which is the wavelength of the Tenoxicam medication.

Antimicrobial susceptibility

Materials were tested for antimicrobial activity using the standard agar-well diffusion method. The test specimens were several human pathogenic bacteria. According to²⁰, the test specimens’ pure cultures were subcultured in nutrient broth medium. Muller–Hinton agar and sterile petri dishes were used to evenly distribute the strain. A sterile cork-borer was used to create a circular well in plates that was 6 mm in diameter. For the purpose of examining antimicrobial activity, a constant weight of 0.5 g was used for each sample, which was pressed into a disc with a diameter of 0.5 cm under a pressure of 50 MPa and subsequently used for the antibacterial resistance test. The plates were then incubated at 37 °C for one day, and the zones of inhibition were determined. Escherichia coli and Klebsiella pneumoniae were used as models for Gram-negative bacteria, and Staphylococcus aureus and Staphylococcus hemolyticus were used as models for Gram-positive bacteria in order to evaluate the antibacterial efficacy of samples (biocomposite) using the agar well diffusion method.

Results and discussion

FTIR of the BG, TXC, and Cs composites before SBF

For BG, C_1, C₂, and C₃ samples: The vibrations of the silicon- oxygen bridges are responsible for the bands at about 463 cm⁻¹ (bending vibrations)²¹. The bands at 560–601 cm⁻¹ correspond to the symmetric stretching of the PO4³⁻ ions²². The TEOS spectrum displays a band centered at about 802 cm⁻¹, assigned to the SiO₄ anti-symmetric stretching modes²³. The band at 1025 cm⁻¹ is caused by asymmetric Si–O–Si stretching of the siloxane bond in the silicate network (Si-O-Si)²⁴. The band at ~ 2349 cm⁻¹ in all samples is due to the asymmetric stretch of gaseous CO₂, which appears and is a very common FT-IR background/residual artifact when the spectrometer sample compartment or beam path is not fully purged or when background subtraction was imperfect²⁵.

For Tenoxicam: The two bands at (1020 and 1088 cm⁻¹) may be due to C–O stretching vibrations consistent with amide functionalities in Tenoxicam²⁶. These two bands (C–O stretching vibrations) overlap with the band at 1025 cm⁻¹, which is due to asymmetric Si–O–Si stretching of the siloxane bond in the silicate network (Si-O-Si) which is confirmed in Fig. 3. The band at 1324 cm⁻¹ is due to sulfonyl (–SO₂–) asymmetric stretching vibrations (S=O) for Tenoxicam²⁷. The band at 1450 cm⁻¹ is due to CH₃ asymmetric deformation (δ_as CH₃) assigned to the presence of methyl in Tenoxicam; this deformation commonly falls around 1450–1470 cm⁻¹²⁸. The 1527 cm⁻¹ peak is a signature of Tenoxicam’s unique fused-ring skeleton. It is assigned to the C = C stretching vibration of the ring system²⁹.

Fig. 3.

Presents FTIR absorption for pure BG, BG loaded with Tenoxicam, chitosan named C1, C2, and C3 samples before immersion in SBF.

The peak at 1635 cm⁻¹ in the infrared (IR) spectrum of Tenoxicam is one of the most important and characteristic bands in the entire spectrum. It is assigned to the C = O stretching vibration (ν(C = O)) of the secondary amide group (–NH–C = O)³⁰. Note that Tenoxicam exists as a zwitterion in the solid state, forming strong intermolecular hydrogen bonds between the amide C = O and the enolic O-H of a neighboring molecule.

This strong hydrogen bonding lowers the stretching frequency of the C = O bond, bringing it down to the lower end of the anticipated range (1635 cm⁻¹), compared to a free, non-hydrogen-bonded C = O (which would be above 1680 cm⁻¹)³¹. Bands around 1900–2200 cm⁻¹ often arise from overtone or combination vibrations of strong fundamental bands in the amide or aromatic regions.

For Tenoxicam, a strong amide I (~ 1635 cm⁻¹) band may have resulted in weak overtones or combination bands in the ~ 1950–2000 cm⁻¹ region. These are not fundamental peaks³². Broad band was noted at ~ 3200–3500 cm⁻¹ due to the existence of O–H stretching (broad hydrogen-bonded OH) and N–H stretching in Tenoxicam³³.

For Chitosan: The band at 877 cm⁻¹ is typically attributed to C–H deformation (out-of-plane bending) of the pyranose ring or glycosidic linkage vibrations³⁴. The broad band from 3000 to 3500 cm⁻¹ in the Chitosan curve is due to O–H stretching vibrations from hydroxyl groups, strongly broadened by hydrogen bonding and often overlaps with N–H stretching vibrations (primary amine groups at C2 in chitosan)³⁴. Analysis of the (C1, C2, and C3) samples revealed a band near 1070 cm⁻¹ corresponding to the symmetric stretching of SiO2 associated with Tenoxicam, which overlaps with the silicate absorption band at 1097 cm^−135,36. The band at 1035 cm⁻¹ increased progressively with the TXC content, as confirmed by deconvolution in the C1, C2, and C3 samples, as demonstrated in Fig. 4.

Fig. 4.

Deconvolution of 900–1250 cm⁻¹ region in C1, C2, and C3 composites.

FTIR of the BG, TXC composites after SBF

In the low-wavenumber region, a band at 460–470 cm⁻¹ was observed in all samples, which is ascribed to the phosphate group’s bending vibrations (PO₄³⁻). The intensity of this band increased due to the presence of phosphate as the dominant functional group in hydroxyapatite, confirming its formation on the sample surfaces³⁷. Additionally, characteristic doublet bands appeared in the 560–600 cm⁻¹ region, corresponding to the bending vibrations of PO₄³⁻. The splitting of this band into two distinct peaks (~ 560 and ~ 600 cm⁻¹) is a well-established feature of hydroxyapatite, reflecting the reduced symmetry of the PO₄³⁻ tetrahedra in the crystal lattice. The presence and enhanced intensity of these phosphate bands further confirm the successful formation and crystallinity of hydroxyapatite in the sol–gel derived samples³⁸.

In the stretching region, a single band was observed at ~ 1040 cm⁻¹; this band was already present before immersing the samples in the SBF solution, where it was previously attributed to Si–O–Si vibrations. After immersion, the intensity of this band increased significantly, which is assigned to the antisymmetric stretching vibrations of the phosphate group (PO₄³⁻). The appearance of this vibration as a single, broad band rather than multiple split peaks is characteristic of sol–gel derived hydroxyapatite, reflecting its nanocrystalline nature and lattice disorder. This feature is commonly reported for apatites synthesized via wet-chemical and sol–gel methods, where reduced crystallinity leads to band broadening and merging in the 1030–1100 cm⁻¹ range³⁸.

It is noteworthy that the band at ~ 1040 cm⁻¹ sharpened following submersion in the SBF solution. This sharpening is ascribed to the release of tenoxicam from the samples into the SBF medium, which reduced the overlapping contributions of the drug and thereby enhanced the definition of the phosphate-related absorption band. An absorption band was observed at ~ 1450 cm⁻¹, which is ascribed to the asymmetric stretching vibrations of carbonate (CO₃²⁻) groups. This band’s existence denotes a partial phosphate substitution (PO₄³⁻) by carbonate in the apatite lattice, known as B-type carbonation, which is the most common in sol–gel derived hydroxyapatite³⁹.

Note that the band at ~ 1450 cm⁻¹ was already present before immersing the samples in SBF and was initially attributed to CH₃. After immersion, however, this band became wider and its intensity increased markedly, indicating an overlapping of CH₃ vibrations with the asymmetric stretching of carbonate (CO₃²⁻). This overlap suggests the incorporation of carbonate into the hydroxyapatite lattice during the hydrolysis process. This study demonstrates that the bioglass/Chitosan system serves as an effective substrate for hydroxyapatite formation, resulting in a notable rise in the synthesized composites’ bioactivity Fig. 5.

Fig. 5.

(FTIR) patterns of BG, TXC, Cs, and TXC-loaded BG/Cs composites after soaking for 33 days in SBF.

XRD results of the (BG)

The Sintered Bioglass’s XRD peak after two hours at 800 °C was enough to prove a typical crystalline structure. A broad hump, which is indicative of particles with an amorphous structure found in silicate glass, emerged at an angle of roughly 23°. Indicating the presence of the calcium hydrogen phosphate hydroxide (Ca₉HPO₄(PO₄)₅OH) crystalline phase, the bands matched the standard density function #00–046-0905. The calcium hydrogen phosphate hydroxide peaks were found at θ2 values of 10.8°, 22.8°, 31.7°, 32.8°, 34°, 39.8°, 46.8°, and 49.4°.

XRD of the BG, tenoxicam, chitosan NPs composites before and after SBF

The X-ray diffraction (XRD) patterns of BG/Cs/TXC composites (C1, C2, and C3) and BG sintered at 850 °C for two hours are displayed in Fig. 6.

Fig. 6.

(XRD) patterns of TXC-loaded BG/Cs composite before immersion in SBF.

At the aforementioned 2θ values, the peak intensities strongly suggested the presence of the calcium hydrogen phosphate hydroxide (Ca₉HPO₄(PO₄)₅OH) crystalline phase, and the angular locations and peak intensities matched the standard likelihood function No. 00–046-0905.

In addition to a constant peak at 20.5° 2θ, two prominent diffraction peaks associated with the TXC phase were observed at 2θ values of 25.8° and 16.7°, with intensity gradually increasing from C1 to C3. Another peak was seen at 37.9° 2θ, with strength gradually increasing from C1 to C3. At 29° 2θ, a clear calcium carbonate (calcite) peak was seen. Furthermore, all samples showed a broad, sharp-apex hump at 25.8° 2θ, which, in accordance with ASTM 075–1381 and JCPDS standards, indicates the presence of silicon dioxide (SiO2) crystals⁴⁰.

The XRD patterns of the BG, C1, C2, and C3 composites following a 33-day immersion in SBF are shown in Fig. 7. The TXC peaks at 20.5° 2θ were seen to fade away after immersion, indicating that the drug Tenoxicam was released from the composites during the SBF soaking operation.

Fig. 7.

(XRD) patterns of TXC-loaded BG/Cs composite after soaking for 33 days in SBF.

The disappearance of the broad hump characteristic of amorphous silica at ~ 23° indicates an active dissolution of the silicate surface layer in the SBF solution. This process leads to the release of silicate (Si⁴⁺) and calcium (Ca²⁺) ions into the surrounding medium. Furthermore, as shown in JCPDS #09–0432, new layers of hydroxyapatite were deposited on the surfaces of the composites after soaking in SBF. The consequent increase in ionic concentration, particularly of calcium and phosphate ions, promotes the nucleation and growth of a new crystalline hydroxyapatite phase. This is evidenced by the emergence of its characteristic diffraction peaks in the XRD pattern (e.g., at 2θ ≈ 25.5° and 29.4°). Therefore, the erosion of the amorphous phase (disappearance of the hump) and the formation of the crystalline hydroxyapatite phase are interconnected manifestations of a single bioactive process. From sample C1 to sample C2, the strength of the hydroxyapatite peaks gradually decreased before increasing in sample C3. The deconvolution analysis of the three strongest hydroxyapatite peaks (211), (300), and (202) formed on the surfaces of BG composites C1, C2, and C3 is shown in Fig. 8⁸.

Fig. 8.

a Deconvolution study of the processed hydroxy apatite’s three strongest peaks on the exterior of BG, C1, C2, and C3 composites: (211), (300), and (202). b The apatite peaks’ crystal size and c their summation crystal size following a 33-day immersion in SBF.

The crystallite size of the apatite peaks was calculated using the well-known Scherrer equation.

1

The crystallite size of the apatite peaks—which were determined by the deconvolution process—was calculated. Where B is the diffraction peak’s full measurement width at half maximum (FWHM), expressed in radians, θ is the diffraction angle (in degrees), λ is the emission wavelength of the X-ray beam (1.5418 Å for Cu Kα rays), Xs is the crystal diameter (in nanometers), and k is the shape factor (valued at 0.94)⁴⁰.

For the C1, C2, and C3 composites, the crystal size for each peak that corresponded to the (211), (300), and (202) planes was identified. It was discovered that C1 was larger than C2, and the aggregate crystal size of these three conspicuous peaks was larger in the C3 sample than in the C1 sample. This suggests a more extensive HA development on the C3 surface, which will be further supported by the SEM and FTIR data.

This formation is characterized by increased porosity and degradation, making it appropriate for drug delivery systems, tissue engineering, and biological applications.

SEM & EDXA results of the BG, TXC, Cs composite

Figure 9 presents the SEM micrographs, which were displayed at 1000× and 5000× magnifications, offering comprehensive information on the surface shape of two samples: the bioglass sample (BG) free Tenoxicam and the 3TXC/BG/Cs composite (C3). A key feature of bone tissue engineering is the porous surface structure of both materials, which has interconnecting open holes that let small blood vessels flow through them to deliver oxygen and nutrients to the inside tissues. The 3TXC/BG/Cs composite (C3) has a larger porosity than the BG alone, according to the comparison of the two materials. The inclusion of chitosan in the composite is responsible for this increase in porosity. The loading capacity of physiological fluids into the biomaterial is improved by higher porosity, which is beneficial⁴¹.

Fig. 9.

Micrographs of a BG and b C3 taken using a scanning electron microscope (SEM). c BG and d C3 EDXA profiles prior to SBF immersion.

The biomaterial’s integration and functionality in biological contexts depend on the interfacial interactions that are subsequently encouraged by this composite between the biomaterial and physiological fluids. The porosity of the composite is critical in bone tissue engineering. It allows for the simple flow of fluids, organic materials, and biological cells through the material, which is necessary for cell proliferation and the production of new bone tissue, particularly osteoblasts.

Furthermore, the Energy Dispersive X-ray Analysis (EDXA) profiles of the BG-free TXC show the presence of silicon (Si), phosphorus (P), and calcium (Ca), the primary ingredients of bioactive glass (BG). In contrast, the carbon content in the 3TXC/BG/Cs composite (C3) is larger due to the carbon poly-chain structure of chitosan, which is absent from the BG-free TXC. This compositional difference highlights the impact of chitosan nanoparticles (Cs) on the composite’s structural and chemical characteristics.

A composite’s capacity to promote the development of a calcium phosphate (CP) coating on its outside after immersion in SBF is commonly used to evaluate its bone bonding capability. Certain materials, conversely, can form a direct link with live bone without displaying a discernible CP layer. After thirty-three days of SBF immersion, SEM micrographs of the surfaces of BG-free TXC and the 3TXC/BG/Cs composite (C3) are displayed in Fig. 10a and b. These pictures show spherical particles on the surfaces, which suggests the creation of CP layers. We can also identify this layer through the ratio of calcium to phosphorus present in the dispersive X-ray analysis.

Fig. 10.

Images from scanning electron microscopy (SEM) of a BG and b C3. EDXA profiles of c BG and d C3 following a 33-day immersion in SBF.

The 3TXC/BG/Cs composite (C3) exhibits more spherical particles (sphere-like crystals) on its surface than the BG sample, according to the SEM micrographs. This would indicate a more widespread production of the hydroxyapatite (HA) layer⁴². Since hydroxyapatite is the fundamental building block of live bone, the more quickly it is precipitated, the better the biocompatible sample³. A HA layer covers the whole surface of the C3 composite because of its increased porosity, but the BG sample exhibits less coverage. EDX spectra of the BG-free TXC and 3TXC/BG/Cs composite (C3) after immersion demonstrate an increase in calcium (Ca) and phosphorus (P) levels and a decrease in silicon (Si) concentration, as presented in Fig. 11c and d. This data supports the SEM findings of HA layer formation. Moreover, the EDX spectra reveal higher concentrations of Ca and P on the C3 surface compared to the BG surface after soaking for 33 days⁴¹. The morphological change of sample C3’s surface before and after immersion in SPF at various magnifications is depicted in Fig. 11a and b, which also displays the spherical forms of hydroxyapatites that are developed on the sample surface.

Fig. 11.

Images from scanning electron microscopy (SEM) of a C3 before to and b following 33 days of immersion in SBF. Comparison of c BG before and after immersion and d C3 before and after immersion using EDXA profiles in SBF for 33 days.

Bioactivity assessment: Ca/P ratio and apatite formation

2

3

After immersion in SBF, the calcium-to-phosphorus ratio of sample C3 was calculated using Eqs. (2,3) from Fig. 11d, c, yielding a Ca/P molar ratio of 1.56. This value falls within the typical range reported for calcium-deficient, bone-like hydroxyapatite (Ca/P ≈ 1.5–1.67), rather than the ideal stoichiometric hydroxyapatite (Ca/P = 1.67), indicating the formation of a bone-like apatite phase on the surface of C3. Similarly, for the BG sample, the Ca/P molar ratio was determined to be 1.63 after SBF immersion. This result also aligns with the range characteristic of bone-like hydroxyapatite, confirming the bioactive nature of the surface and the development of an apatite layer consistent with bone mineralization processes.

Drug (Tenoxicam) release

The UV and visible absorption spectra of Tenoxicam are displayed in Fig. 12. When the strength of Tenoxicam’s absorption peak was measured, it was discovered to be at 370 nm. The Tenoxicam solution’s absorption peaks were observed at the same wavelength in the other research⁸. Tenoxicam concentrations were prepared from the basic solution portions diluted with distilled water at pH 7.4 Table 2^43,44.

Fig. 12.

a UV–VIS absorption of Tenoxicam concentration, b calibration curve of the released drug Tenoxicam.

Table 2.

Standard absorption values of Tenoxicam in distilled water, pH = 7.4, with different concentrations.

No	Concentration (mg/ml)	Absorbance (%)
1	0.0682	2.649
2	0.0568	2.207
3	0.0454	1.764
4	0.0341	1.325
5	0.0227	0.882

Figure 13 shows the demonstrations of the three-region cumulative release curve of Tenoxicam encapsulated in chitosan polymer with bioglass in the in vitro test. Three phases were utilized to investigate the profile of medication release: phase I involved an initial increasing release (within 24 to 96 h), phase II involved a steady release (within 4 to 16 days), and phase III involved a slow release (from 16 to 33 days). Phase (I) saw 65.00%, 60.00%, and 54.92% of C1, C2, and C3 releases, respectively. The release curves indicate a decreasing release rate between 4- and 16-day phase (II). The corresponding release percentages were 75.60%, 77.16%, and 73.19%. The drug produced from compounds C1, C2, and C3 phase (III) was steady for 24 days after 16 days, reaching 87.46%, 89.53%, and 85.40%, respectively. For the three concentrations C1, C2, and C3, the profile is comparable, as would be expected from bioglass/chitosan composites, which support the medication⁴⁵.

Fig. 13.

Tenoxicam absorbance (a.u.) at 370 λ during soaking from start to (33) days.

Diffusion and polymer breakdown are two variables that affect medication release. Based on the release profiles, it seems that the interaction between the drug’s NH and the OH of the chitosan polymer causes the release mechanism of Tenoxicam to be mostly diffusion, followed by polymer degradation. According to the experimental findings, Bioglass (C2) released Tenoxicam more quickly than Bioglass/Chitosan composites in sample (C1)⁴⁶. This is because, whereas C1 and C3 may be supported by diffusion based on the chemical interaction of the amino groups of the chitosan polymer and the carbonyl groups of the Tenoxicam drug, the release of the drug from Bioglass (C2) may be more influenced by the degradation of the encapsulated polymer in the latter period. Moreover, by fitting the release data to several mathematical equation models, the release kinetics of Tenoxicam from compounds C1, C2, and C3 were ascertained, as shown in Table 3.

Table 3.

Tenoxicam medication release percentage from compound samples.

Sample time (d)	C1		C2		C3
	mg/ml	%	mg/ml	%	mg/ml	%
At start	0	0.00	0	0.00	0	0.00
0.04167	3.294889523	5.37	3.320630848	5.491644	0.538831	6.52037
0.083	5.302712827	14.02	4.350283824	11.11649	0.479984	12.32863
0.125	4.144353229	20.78	5.251230178	17.80092	0.55004	18.98464
0.25	5.071040907	29.05	6.769968318	24.10995	0.580064	26.00397
0.5	4.581955743	36.52	6.615520371	30.85493	0.58807	33.12019
1	6.769968318	47.57	6.4353311	37.90264	0.706164	41.66546
2	5.766056666	56.97	6.718485669	45.05934	0.556845	48.40382
4	5.096782231	65.28	5.688832693	56.2061	0.872297	58.95945
8	3.114700253	70.36	5.817539315	65.88762	0.746997	67.99884
16	3.475078794	76.03	5.508643422	75.47832	0.80024	77.68251
24	6.641261696	86.86	8.391671755	86.53427	0.886308	88.40769
33	8.057034538	100.00	8.674826323	100	0.957966	100
Total amount	61.31583472		73.54296381		67.05615007
t value	−5.78187		−5.22974		−5.42806
p value	< 0.00001		0.000012		< 0.00001

*P value < 0.01 was considered statistically significant.

From Table 4, evidently, the release kinetics of C1, C2, and C3 followed the Zero-order Release model, with the release kinetics parameters of different Tenoxicam-loaded BG/Cs composites.

Table 4.

Release kinetics parameters of different TXC-loaded BG/Cs composite.

Sample	R2 value			Zero-order release			0k	D
	Korsmeyer–Peppas model	Higuchi release model	Hixson Crowell release model	R2 value	t50* (hours)	t90** (hours)
C1	0.3965	0.5669	0.9812	0.9938	6.363	11.453	7.8581	8.0609
C2	0.4181	0.557	0.9737	0.9867	13.181	23.726	7.5866	8.6779
C3	0.987	0.9812	0.9523	0.9693	22.837	41.106	6.5684	9.5930

R²-value is the regression coefficient value, ₀k = Zero-order release rate constant, D is the total dose or total amount of drug in the dosage form sample, t₅₀* is the time needed for 50% of the medication to be released, t90** is the time needed for 90% of the drug to be released, and so on. (IBM SPSS statistical program, version 30.1).

Zero-order kinetics of drug release indicates that: The rate of drug release remains constant across time. This has the advantage of maintaining a steady plasma concentration in the blood for an extended period of time. For medications that need a steady dosage to be effective and prevent swings, this is crucial. This is what the drug TXC used in the samples under study achieved. Figure 14, expressed as a percentage, displays the distribution of drug profile release across the whole implantation time.

Fig. 14.

a The release amount of TXC concentration from samples for (1–33) days, b release profile of Tenoxicam in terms of the percentage (%) of TXC released as a function of time.

Antimicrobial susceptibility

Resistance to antimicrobials, which harms human health, increases the risk of infections and increases the death rate from serious, life-threatening illnesses, is currently the biggest health issue in every country on the globe. As a result, we prefer to include a chemical that makes the organic material utilized automatically resistant to many forms of germs. This strengthens its position as an alternative organic material in post-surgical implants while also providing the essential resistance to the majority of germs tested on it. The agar-well-based diffusion technique was used to gauge how antibacterial effectiveness differs among different microorganisms, such as Staphylococcus aureus and Staphylococcus hemolyticus as models for Gram-positive bacteria and Escherichia coli and Klebsiella pneumoniae as models for Gram-negative bacteria.

The antibacterial efficacy of BG and TXC-loaded BG/Cs composites against four distinct bacterial strains is depicted in Fig. 15; Table 5. The diameters of inhibition zones of C1, C2, and C3 were (35 ± 0.7), (22 ± 0.2), and (29 ± 0.6) mm, respectively, when exposed to (Staphylococcus aureus). While the sample BG-free of the drug Tenoxicam did not show any response or inhibition zone around the sample when tested with this microbe.

Fig. 15.

Microbial sensitivity of BG, C1, C2, and C3 against pathogenic bacteria was tested.

Table 5.

Microbial susceptibility of BG, C1, C2, and C3 to test harmful microorganisms.

No.	Isolate name	Diameter of Inhibition zone (mm)
		Samples
		BG	C1	C2	C3
1	Klebsiella pneumoniae	23 ± 0.6	44 ± 0.8	39 ± 0.9	35 ± 0.9
2	Escherichia coli	35 ± 0.8	39 ± 0.6	39 ± 0.5	28 ± 0.3
3	Staphylococcus aureus	0 ± 0	35 ± 0.7	22 ± 0.2	29 ± 0.6
4	Staphylococcus haemolyticus	14 ± 0.8	18 ± 0.6	36 ± 0.8	25 ± 0.2

While after exposure to (Staphylococcus haemolyticus) of BG, C1, C2, and C3; they were (14 ± 0.8), (18 ± 0.6), (36 ± 0.8), and (25 ± 0.2) mm, respectively. While they demonstrated diameters of inhibition zones of (35 ± 0.8), (39 ± 0.6), (39 ± 0.5), and (28 ± 0.3) mm, respectively, when exposed to (Escherichia coli).

Finally, as seen in Fig. 16, they were (23 ± 0.6), (44 ± 0.8), (39 ± 0.9), and (35 ± 0.9) mm, respectively, following exposure to Klebsiella pneumoniae. According to these findings, BG and TXC-loaded BG/Cs composites have antibacterial properties, which could result from the composites’ capacity to release TXC ions^47,48. In contrast to BG, C1, and C2, C3 had stronger antibacterial activity effects. This suggests that any infection that may develop during the implantation process can be treated with BG and the composites created.

Fig. 16.

Microbial sensitivity of BG, C1, C2, and C3 against pathogenic microorganisms.

By calculating the cumulative widths of the inhibition zones for all four species tested, we can distinguish between the samples, and it is obvious in Fig. 17 that sample C3 produced the best results for all organisms tested, as compared with the other samples, C1, C2, and BG.

Fig. 17.

Microbial sensitivity of BG, C1, C2, and C3 against pathogenic microorganisms.

Conclusions

This study aimed to develop a bioactive composite of bioglass (BG) and chitosan (Cs) loaded with the anti-inflammatory drug Tenoxicam (tenoxicam) for bone tissue engineering. Tenoxicam composites with weight percentages of 1, 2, and 3 wt% were prepared using the sol-gel method, ensuring high porosity and bioactivity. Discs were fabricated and characterized by FTIR, XRD, scanning electron microscope, and EDX. Antibacterial activity was tested against Gram-positive and Gram-negative bacteria using the agar diffusion method. Drug release behavior was monitored for 33 days in simulated body fluid (SBF) using UV spectroscopy at 370 nm.

FTIR and XRD results confirmed the formation of hydroxyapatite (HA) on the composite surface after SBF immersion, indicating strong bioactivity. FTIR spectra revealed characteristic phosphate and silicate peaks, supporting the integration of bioglass and chitosan. SEM images showed a porous structure, particularly in samples with higher Tenoxicam content, promoting fluid penetration and cell adhesion. EDX analysis detected increased calcium and phosphorus after soaking, confirming HA deposition. Antimicrobial studies showed inhibition zones against Staphylococcus aureus, Staphylococcus hemolyticus, Escherichia coli, and Klebsiella pneumoniae. Drug release profiles demonstrated sustained Tenoxicam delivery, with C3 (3 wt%) exhibiting the highest release.

Although this study demonstrates successful fabrication, in vitro bioactivity, antibacterial performance, and sustained tenoxicam release under the selected condition, further work is needed to strengthen translational relevance. Future studies will include in vitro cytocompatibility testing, pH-dependent drug release under clinically relevant environments, quantitative ion-release and pH monitoring during immersion/release, BET surface area/porosity analysis to correlate microstructure with release behavior, and expanded antimicrobial evaluation including antifungal activity, followed by in vivo validation.

Author contributions

Conceptualization, M.E, A.A, A.M; Data curation, M.E, A.A, N.E, A.Z, A.M; Formal analysis, M.E, A.A, A.A, A.E, M.M, N.E, A.Z, A.M,; Investigation, A.E, M.M, N.E, A.Z, A.M; Methodology, Writing – original draft, M.E, A.A, A.A, A.E, A.M; Writing – review & editing, M.E, A.A, A.A, A.E, M.M, N.E, A.Z, A.M. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported and funded by the Deanship of Scientific Research at Imam Mohammad Ibn Saud Islamic University (IMSIU) (grant number IMSIU-DDRSP2601).

Data availability

The data presented in this study are available on request from the corresponding author.

Declarations

Competing interests

The authors declare no competing interests.

Consent for publication

Not applicable.

Clinical trial number

Not applicable.

Ethics declaration

Not applicable.