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Article

Enhancing PEEK Surface Bioactivity Through Phosphate and Calcium Ion Functionalization

by
Lillian V. Tapia-Lopez
1,*,
Antonia Luna-Velasco
2,
Carlos A. Martínez-Pérez
3,
Simón Yobanny Reyes-López
4 and
Javier S. Castro-Carmona
3,*
1
Department of Chemistry and Biochemistry, University of Texas at El Paso (UTEP), El Paso, TX 79968, USA
2
Department of Environmental Sciences, Advanced Materials Research Center (CIMAV), Miguel de Cervantes Saavedra 120, Industrial Complex Chihuahua, Chihuahua 31136, Mexico
3
Institute of Engineering and Technology (IIT), Autonomous University of Juárez City (UACJ), Av. del Charro 450, Partido Romero, Ciudad Juárez 32310, Mexico
4
Institute of Biomedical Sciences (ICB), Autonomous University of Juárez City (UACJ), Av. del Charro 450, Partido Romero, Ciudad Juárez 32310, Mexico
*
Authors to whom correspondence should be addressed.
Coatings 2025, 15(11), 1359; https://doi.org/10.3390/coatings15111359
Submission received: 24 October 2025 / Revised: 13 November 2025 / Accepted: 17 November 2025 / Published: 20 November 2025
(This article belongs to the Special Issue Surface Coating, Functionalization, and Characterization of Cell- and Tissue-Based Biomaterials)
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Abstract

Inert polymeric implants must evolve to enhance their biological interactions with host tissue, triggering positive cellular responses and promoting tissue bonding and integration. Poly-ether-ether-ketone (PEEK) is widely used as an implant material; however, its inert nature results in limited biological interactions. Various surface modification techniques have been investigated to enhance its bioactivity and overall biological performance. In this study, the PEEK surface was bioactivated through a chemical treatment involving two steps: surface activation using low-pressure oxygen plasma, followed by biofunctionalization with phosphate and calcium ions. Comprehensive surface characterization by contact angle, scanning electron microscopy (SEM), X-ray photoelectron spectroscopy (XPS), and Fourier-transform infrared (FT-IR) confirmed the effect of plasma and the ionic surface incorporation. The biological response was evaluated through cell viability, adhesion, and proliferation in NIH/3T3 fibroblasts and HOS osteoblasts, and the results indicated the efficacy of the surface modifications. Therefore, the proposed treatments provide an efficient strategy to improve the biological performance of PEEK-based implants.

1. Introduction

The manufacturing of personalized implants has been made possible by 3D printing [1,2]. This printing is an excellent achievement in medical areas such as orthopedics, maxillofacial, cranial, and spine, where implants can vary between patients due to bone defects and differences in physical complexion. This type of impression aligns with appropriate materials that meet mechanical, chemical, and biological requirements. Polyether ether ketone (PEEK) is a thermoplastic polymer widely used in trauma and spinal fixation [3,4] and is considered a metal replacement material for its mechanical properties, biocompatibility, and radiolucency [4,5], making it appropriate for bone implantation and imaging processing [5,6]. The tensile strength of PEEK ranges from 90 to 100 MPa [3,7], and the elastic modulus of around 3.6 GPa [3,5], indicating its stiffness is more like bone compared to metals [4,8]. In addition, PEEK exhibits wear and abrasion resistance, high-temperature performance, dimensional stability [3], and creep resistance [4,9]. Also, this material is chemically stable, low or non-toxic, non-inflammatory [3], and shows no mutagenic or carcinogenic effects [9]. However, like many biomaterials, PEEK has low surface energy (hydrophobic nature) and is considered bioinert [9,10]. In fact, evidence from proteomic studies has suggested that PEEK may alter certain types of proteins, which have been related to low cell proliferation on its surface [11].
Current research is focused on the bioactivity of biomaterials due to the significance of surface properties in influencing cell behavior [12,13]. These surfaces are arising as promising candidates for suitable implant materials because they mimic the natural extracellular matrix (ECM) properties, creating an environment that promotes a favorable cell response [14]. Modifying the biomaterial surfaces by enhancing hydrophilicity, surface energy, and chemistry can improve their adhesive properties [14,15,16] and bioactivate their surfaces for better implant tissue interaction. In addition, bioactivity may reduce the healing period [17].
Previous work from our group demonstrated that coating zirconia surfaces with laminin-5 protein significantly improved cellular response [18]. Moreover, because ECM proteins interact with cells mainly through the RGD peptide sequence (Arg–Gly–Asp) [14], we have also verified that direct immobilization of RGD on polydopamine-coated PEEK enhanced cell attachment while avoiding protein instability [19].
Similarly, calcium phosphate treatments have demonstrated an effective cellular response, facilitating the osteoblastic differentiation into mature osteoblasts (OBs) able to synthesize bone matrix and contribute to bone mineralization [20,21]. For instance, biomaterials coated with calcium phosphate (CaP) have shown favorable osseointegration [22,23]. However, other authors have reported that coatings with hydroxyapatite or octacalcium phosphate, although beneficial, can detach over time due to weak physical bonding, and the resulting particle debris may compromise implant stability [24]. In addition to bioactive coatings, various surface activation methods have been explored to enhance polymer bioactivity and promote calcium phosphate deposition. Electrodeposition enables rapid, controllable mineralization of polymer scaffolds, followed by immersion in simulated body fluid (SBF) to enhance apatite formation [25]. Laser activation has also been employed to induce selective mineralization on PLLA films, facilitating apatite nucleation after subsequent SBF incubation [26]. Furthermore, chemical etching and functionalization have been employed to introduce reactive groups that promote Ca/P nucleation on polymeric scaffolds, leading to the formation of uniform hydroxyapatite-like coatings after immersion in SBF [27].
While the approaches described above aim to impart bioactivity to otherwise inert materials, there also exist intrinsically bioactive materials, such as bioceramics, in which calcium and phosphate are part of their bulk composition. Hydroxyapatite, tricalcium phosphate, and bioactive glasses are widely used because they bond directly to bone and support both osteoconduction and osteogenesis. Nonetheless, several reviews have noted that their main mechanical drawback is that they are brittle, exhibit low fracture toughness, and that their strength decreases further when high porosity is required for bone ingrowth, which limits their application in load-bearing sites [28,29,30].
In this context, our approach aims to combine the mechanical advantages of PEEK with the bioactivity of calcium and phosphate ions, thereby bridging the gap between bioinert and bioactive systems. Unlike conventional coating methods, this process does not create a deposited layer but rather achieves surface functionalization through ionic immobilization. Thus, PEEK surfaces were therefore activated by low-pressure oxygen plasma and subsequently functionalized with phosphate and calcium ions to promote bioactivity. Plasma treatment increases surface energy and introduces oxygen-containing groups, enabling further ion modifications [31,32]. To evaluate the modified material, in vitro experiments were conducted using the cell lines NIH/3T3 mouse fibroblast and HOS human osteoblast cell lines to determine cell viability, adhesion, spreading, and proliferation on PEEK surfaces. The results demonstrated the feasibility of these surface modifications to enhance the bioactivity of PEEK and, consequently, the cellular response.

2. Materials and Methods

2.1. Materials

Commercial-grade PEEK was obtained from Cera Direct (Shenzhen, China). Calcium hydroxide and sodium dihydrogen phosphate monohydrate were purchased from Sigma-Aldrich (St. Louis, MO, USA). Cell culture reagents, including DMEM-F12, Trypsin-EDTA, and Fetal Bovine Serum (FBS), were supplied by Gibco, Thermo Fisher Scientific (Waltham, MA, USA). For cell staining, Hoechst 33342 and Phalloidin CF568 were purchased from Abcam (Cambridge, UK) and Biotium (Fremont, CA, USA), respectively.

2.2. Sample Fabrication and Surface Modification

2.2.1. Fabrication and Polishing

PEEK disc-shaped samples were digitally designed using the SolidWorks© program 2021 and machined in a Roland DWX-51D (Hamamatsu, Japan). Each disc was 3 mm thick, with diameters of 18 mm (contact angle), 5.8 mm (cell viability), and 10 mm (other analysis). Samples were sequentially polished using a GPX 200 Leco brand micro-rotator (Mumbai, India), with 600- and 1200-grit alumina papers for 20 s each. Samples were cleaned by ultrasonic agitation in ethanol (30 min) and DI water (5 min). The resulting polished specimens were referred to as control samples.

2.2.2. Surface Activation

The polished samples were treated with oxygen plasma using a low-pressure Diener Plasma Technology system (Ebhausen, Germany). During treatment, the gas flow was kept at 5 sccm, with a chamber pressure of about 0.5 mbar, a power supply of 50 W, and a frequency of 40 kHz. The vacuum pump operated at 1.5 m3/h. After 5 min of plasma exposure, the samples were rinsed with deionized water to stabilize them [33]. Plasma treatment enhances hydrophilicity and generates oxygen-containing groups [16,34] that serve as anchoring sites for subsequent ionic functionalization [31]. These samples were labeled PL-S.

2.2.3. Surface Functionalization

In the functionalization process, some samples were treated with calcium, and others were exposed to both phosphate and calcium ions. For calcium incorporation, plasma-treated samples (PL-S) were immersed in an aqueous solution of 30 mM Ca(OH)2 under continuous stirring at 40 °C for 1 h (Ca-S samples). For the samples functionalized with phosphate and calcium ions, PL-S samples were first immersed in an aqueous solution of 50 mM NaH2PO4·H2O at 80 °C for 2 h. Then, 30 mM Ca(OH)2 was added to the solution, and the reaction continued for one more hour under magnetic stirring. After each functionalization, the samples were rinsed and then dried (PCa-S samples).

2.3. Surface Evaluation

2.3.1. Water Contact Angle Measurement

Static water contact angles were determined using a First Ten Angstroms 200 instrument (FTA-200, Newark, CA, USA) by depositing a 2 µL DI water droplet on different surface points. Measurements were taken 20 min after each surface modification; values represent the mean ± SD (1.72° to 6.29°).

2.3.2. Crystallinity Analysis

Fourier-transform infrared spectroscopy with attenuated total reflectance (FTIR–ATR) was performed using a Perkin Elmer Spectrum GX system (Waltham, MA, USA) to evaluate the crystallinity of plasma-treated samples (PL-S) compared with the control sample in the 1500–1200 cm−1 region. Crystallinity was evaluated based on the absorption bands at wave numbers 1305 and 1280 cm−1, as these peaks are reported for crystallinity, and the ratio of 1305/1280 cm−1 represents the crystallinity index (CI) [8]. The percentage of surface crystallinity was calculated following the ASTM F2778-09 standard [35] using the equation: Crystallinity [%] = (CI − 0.728)/1.549 × 100.

2.3.3. Surface Topography Analysis

The topographical analysis was performed using a JEOL7401 JSM scanning electron microscope (SEM, Tokyo, Japan). The parameters were set at 10 kX for magnification, 2 kV for the acceleration voltage, and 8 mm for the working distance. The field emission electron gun utilized a tungsten tip emitter operating under ultra-high vacuum conditions.

2.3.4. Chemical Composition Analysis

Surface chemical composition was analyzed by X-ray photoelectron spectroscopy (XPS) using a Thermo Scientific Escalab 250xi system (Waltham, MA, USA) equipped with a monochromatic Al Kα source (hν = 1486.86 eV). The samples were studied 48 h after each treatment using a 10 eV electron gun voltage, a 650 µm spot size, and a 1 eV energy step.

2.4. Cell Culture Assays

Cell culture procedures previously established and validated in our laboratory [19] were used for all experiments. Cells were maintained in DMEM-F12 supplemented with 5% FBS and 1% streptomycin at 37 °C, 5% CO2, in a humidified incubator. Trypsin-EDTA was used for cell detachment during subculturing, which was performed every three days.

2.4.1. Cell Viability Analysis

For the MTT assay [36], samples were placed in 96-well plates, and fibroblasts and osteoblasts were seeded at a density of 2 × 104 cells using 200 µL of DMEM-F12 supplemented with 5% FBS. The plates were incubated at 37 °C in a humidified atmosphere containing 5% CO2 for 24 h. Thereafter, the media were replaced with DMEM containing 0.5 mg/mL MTT and incubated for another 4 h period. The medium in each well was discarded and rinsed. The formazan crystals produced during the MTT reaction were dissolved with dimethyl sulfoxide, and the absorbance was read at 570 nm using a Varioskan Lux microplate reader from Thermo Fisher (Waltham, MA, USA). Results were expressed as optical density (OD) relative to control wells. Three independent experiments were performed (n = 3).

2.4.2. Cell Proliferation Analysis

Samples were placed in 24-well plates, and fibroblast and osteoblast cells were seeded at a density of 2 × 104 cells for 24 and 96 h, under the same incubation conditions described previously. Cells were fixed with 3.75% paraformaldehyde (15 min, 4 °C), permeabilized with 0.2% Triton X-100 (10 min, 25 °C), blocked with bovine milk (1 h, 25 °C), and cell nuclei stained with Hoechst (30 min, 37 °C). Samples were rinsed with PBS between steps, imaged using a Zeiss LSM-700 confocal microscope (Jena, Germany), and counted using ImageJ software (version 1.53t). At least 20 images per sample were taken for each sample group (n = 2).

2.4.3. Cell Adhesion and Spreading Analysis

Samples were placed in 24-well plates, and fibroblast and osteoblast cells were seeded at a density of 2 × 104 cells for 24 h, under the same incubation conditions described previously. After fixation, permeabilization, and blocking, actin filaments were stained with Phalloidin CF568 (20 min, 25 °C), and nuclei with Hoechst 33342 (30 min, 37 °C). Samples were rinsed with PBS between steps. Fluorescent images were obtained at 40× magnification using a Zeiss LSM-700 confocal microscope (Jena, Germany).

2.5. Statistical Evaluation

Data are expressed as mean ± standard deviation (SD). Statistical analysis was conducted with GraphPad Prism v9 using one-way ANOVA. A statistically significant value was considered for p < 0.05.

3. Results

As previously mentioned, three different surface treatments were applied to PEEK samples, along with one untreated sample used as a control. These samples underwent a series of analyses and experiments, and the results were compared to assess the effects of each surface treatment. Table 1 summarizes the sample types and their corresponding labels to facilitate understanding of the results presented.

3.1. Hydrophilicity

The contact angle results for all PEEK sample surfaces are presented in Figure 1.
The control sample is the least hydrophilic surface with an angle of 86.49° ± 1.73. After oxygen plasma treatment, the PL-S sample showed a high decrease in the contact angle value, 25.98° ± 1.35. The functionalization with calcium ions, Ca-S sample, showed an angle of 58.42° ± 6.29, and the functionalization with phosphate and calcium ions, PCa-S sample, had a value of 43.03° ± 6.00. The functionalized surfaces had a higher standard deviation, indicating that the surface energy varies across different parts of the surfaces, probably due to the heterogeneity of ion distribution.

3.2. Crystallinity

Figure 2 shows the spectra of the control and oxygen plasma-treated (PL-S) samples. The characteristic peaks at 1305 and 1280 cm−1, corresponding to the crystalline phase of PEEK [8], appear in both spectra without any noticeable variation after treatment.

3.3. Topography

Topographical changes were observed through SEM micrographs among the treated samples (Figure 3). The control sample’s surface (Figure 3a) showed a smooth morphology with few irregularities. In contrast, the plasma-treated sample (PL-S, Figure 3b) exhibited mild surface etching in localized areas. Functionalization with calcium ions (Ca-S, Figure 3c) produced scattered surface features. In the same way, the phosphate–calcium functionalized sample (PCa-S, Figure 3d) displayed a similar topographic pattern, with granular artifacts distributed over the entire surface.

3.4. Chemical Composition

XPS analysis confirmed notable differences in elemental composition among the PEEK surfaces. Table 2 summarizes the percentage composition of each sample, and Figure 4 displays their survey spectrum and deconvoluted peaks.
The percentage of chemical composition between the control and PL-S samples varies in the amount of oxygen detected over the material surface. After plasma treatment, the PL-S sample showed 26.73 ± 3.62% oxygen compared with 20.54 ± 0.11% from the control, indicating a higher surface oxygen concentration (Table 1). The increased amount of oxygen is due to the formation of oxygen-containing groups on the polymer surface. After ionic treatment, calcium was detected in the Ca-S sample (2.38 ± 1.12% Ca). For the PCa-S sample, both phosphorus and calcium were identified, with an increased calcium concentration (8.20 ± 1.07%) compared with Ca-S (Table 1). This suggests that phosphate incorporation promotes additional calcium binding.
For the XPS survey spectrum, Figure 4a displays the characteristic peaks of every surface. The control and plasma-treated PEEK samples (PL-S) exhibited C1s around 289 eV and O1s around 537 eV peaks. In contrast, calcium-functionalized surfaces (Ca-S) presented an additional Ca2p peak around 346 eV, and the phosphate-calcium modified samples (PCa-S) exhibited both P2p around 133 eV and Ca2p around 346 eV signals.
The peak representations shown in Figure 4b,c were processed using OriginPro 2018, and the binding energies were identified according to the NIST XPS spectroscopy database. For the Ca-S sample, the Ca2p signal was deconvoluted into two peaks. Ca-O was detected at 347.4 and 351.05 eV. For the PCa-S sample, the P2p signal was detected at 133.30 eV, corresponding to CaHPO4, while Ca2p deconvolution resulted in a signal around 347.25 eV for CaHPO4 and Ca-O formation at 347.25 and 350.8 eV.

3.5. Cellular Response

3.5.1. Cell Viability

The optical density (OD) values obtained from the MTT assay after 24 h are shown in Figure 5 for fibroblast and osteoblast cells.
All treated samples exhibited a statistically significant cell viability compared with the control, indicating an improvement in the metabolic activity in response to surface modification. Fibroblasts exhibited a gradual increase in viability across the treatments (Figure 5a), while osteoblasts showed similarly high levels among all plasma-treated and functionalized samples (Figure 5b).

3.5.2. Cell Proliferation

Cell proliferation is shown in Figure 6 for all surface conditions relative to the control.
Both cell lines showed proliferation after 24 h, with a more pronounced increase at 96 h for the plasma-treated and ion-functionalized samples. Fibroblasts exhibited the highest proliferation on the phosphate and calcium ion functionalized surfaces (PCa-S) after 24 h, while at 96 h, proliferation values were comparable among plasma-treated and functionalized samples. Osteoblasts had a clear preference for ion-functionalized surfaces, particularly for PCa-S, at both 24 and 96 h, and the low standard deviation values indicated consistent cellular behavior across replicates. These results confirm the positive influence of surface modification on sustained cell proliferation.

3.5.3. Cell Adhesion and Spreading

Representative fluorescence images of fibroblast and osteoblast cells after 24 h are shown in Figure 7.
Cell adhesion was observed on all surfaces, including the control sample, for both cell types. However, plasma-treated (PL-S) and functionalized (Ca-S and PCa-S) samples exhibited more extended and well-spread morphologies.

4. Discussion

The lower contact angle value observed for the PL-S sample confirms the successful incorporation of oxygen-containing groups onto the PEEK surface during plasma exposure, as reported previously [16,31,34]. Plasma contains ions, electrons, free radicals, and photons generated by ultraviolet radiation. This radiation generates enough energy to break chemical bonds on the surface of the material, and then oxygen binds to reactive sites, creating polar groups (–OH, –COOH, –C=O) [37,38]. These polar functionalities increase surface energy and hydrophilicity. Similar mechanisms have been reported by other authors [39,40], who demonstrated that the extent of oxidation and surface activation depends strongly on plasma parameters, including species flux, exposure time, and gas composition. Following this activation step, ion-functionalized samples exhibited an increase in contact angle compared with the plasma-treated surface, most likely due to changes in surface chemistry and partial charge neutralization of oxygen-containing functional groups. The higher standard deviations observed are likely related to surface heterogeneity arising from non-uniform ion distribution. Overall, the increase in hydrophilicity following plasma and ionic modification relative to the control indicates surface activation, which is expected to promote enhanced cellular response. The FTIR-ATR results indicated a crystallinity index value of around 1.00 for the Control and PL-S sample, with a percentage of crystallinity of 17.88% and 18.03% respectively, suggesting that the oxygen plasma process caused only minor structural changes at the surface level [8]. Although this technique is not a fully quantitative method for determining crystallinity, it offers a comparative indication of surface structural changes. The plasma process used in this study is a low-pressure, non-thermal treatment in which energetic species interact primarily with the outermost molecular layers of the polymer. In this regime, the kinetic energy of the ions and radicals, reported by Versel et al. to be between 6 and 12 eV, is insufficient to penetrate beyond the near-surface region. This behavior is consistent with previous studies reporting that plasma activation modifies the surface chemistry while preserving the bulk properties of PEEK [8,37,41]. In the SEM analysis, the mild etching observed in some areas of the PL-S sample (Figure 3b) compared with the control (Figure 3a) revealed the effect of oxygen plasma on the surface of the polymeric material after 5 min of exposure. In addition, the functionalization also generated morphological changes. Both Ca-S and PCa-S samples exhibited scattered surface features (Figure 3c,d), suggesting that phosphate and calcium ions interacted with the oxygenated surface groups formed during plasma treatment. This was corroborated not only by the contact angle results but also by XPS. The amount of oxygen detected after plasma treatment (Table 2, Figure 4a) confirmed the formation of oxygen-containing groups on the polymer surface [31,32]. Building on this activated surface, in the Ca-S sample, calcium interacted mainly with the oxygenated groups. Under the alkaline conditions of the Ca(OH)2 treatment, these oxygen atoms become partially deprotonated, acquiring negative charges that enable coordination with calcium ions to form stable Ca–O bonds. Similar Ca2+–oxygen coordination has been reported for carboxylate ligands [42]. In our Ca-only sample, this mechanism explains the Ca–O binding observed by XPS, Figure 4b. In the PCa-S sample, the interaction between calcium and phosphate species is strongly influenced by the protonation state of phosphate groups, which depends on the pH of the medium. During the first step with sodium dihydrogen phosphate (NaH2PO4, pH ≈ 5), the main phosphate form is H2PO4−, which interacts with the oxygenated groups generated during plasma activation through hydrogen bonding and electrostatic attraction. When calcium hydroxide is added, the pH rises and the phosphate species partly deprotonate to HPO42−. Under these alkaline conditions, calcium associates not only with phosphate oxygen, forming CaHPO4, but also with oxygenated surface sites on PEEK, leading to additional Ca–O bonds observed in the XPS spectra, Figure 4c. These interactions likely involve both electrostatic attraction and coordination bonding. This behavior is consistent with Chen et al. (2022), who reported that deprotonated phosphate groups coordinate Ca2+ through charge-transfer bonding under alkaline conditions [43].
Hence, by increasing surface hydrophilicity and incorporating bioactive ions, these surface modifications promote protein adsorption and induce conformational rearrangements that expose integrin-binding domains [44], thereby activating intracellular signaling pathways involved in cell adhesion and spreading. Moreover, calcium acts as a signaling messenger that drives osteoblast differentiation and matrix mineralization. Through nitric oxide generation and activation of ERK1/2 and PI3K/Akt pathways, it supports cell growth and survival [20]. The influx of calcium through Ca2+- and K+-regulated channels activates intracellular cascades—CaMKII, calcineurin/NFAT, and CREB—that control osteogenic gene expression and drive matrix mineralization [45]. In addition, phosphate ions contribute to osteoblast differentiation and bone matrix formation by activating pathways such as IGF-1 and ERK1/2, and by enhancing bone morphogenetic protein (BMP) signaling. They also help regulate bone resorption by balancing RANKL and OPG signaling [20]. According to Khoshniat et al. (2011), phosphate-induced activation of the ERK1/2 pathway occurs only in the presence of calcium ions, since both species must reach a threshold concentration to form nanoscale calcium phosphate complexes at the cell interface [46]. Consistently, Vermeulen et al. (2022) reported that biomaterials that release Ca2+ and phosphate ions can modulate several osteogenic signaling cascades, including the ERK1/2, BMP, and Wnt pathways [47]. Both participate in the bone repair process, with osteoinductive and osteoconductive capacities that support protein adsorption, cell adhesion, and new bone growth [20].
Given that calcium and phosphate ions regulate osteoblast activity and survival, cell viability was assessed for both cell types after 24 h (Figure 5). Viable cells were observed in all samples, with higher levels in those treated with plasma and ion functionalization, suggesting that improved hydrophilicity and ionic chemistry favored early cellular attachment and metabolic activity. When focusing on fibroblasts, the viability and proliferation results at 24 h for the plasma-treated samples were inconclusive. At that time, the specific effects of the plasma treatment may have enhanced cell metabolism without promoting division, resulting in metabolically active but non-dividing cells under the evaluated conditions. Nevertheless, fibroblast proliferation values were comparable across all plasma-treated and ion-functionalized surfaces at 96 h, indicating that fibroblast growth was well supported over time, regardless of the applied modification. In contrast, osteoblasts displayed a more consistent response on the phosphate–calcium surfaces (Figure 6), suggesting that this modification provides a more stable and favorable microenvironment for osteogenic activity [17,23].
Finally, enhanced adhesion and spreading observed on the modified samples compared with the control could reflect an improved cytoskeletal organization and focal contact formation, leading to a more effective anchorage of both fibroblast and osteoblast cells. These results demonstrate that hydrophilicity and bioactive ionic species together enhance biocompatibility and cellular interactions at the material interface.

5. Conclusions

This study evaluated the effect of plasma activation and ionic functionalization on PEEK surfaces to improve their biological performance. The effect of such surface modifications was confirmed using several characterization techniques. The contact angle verified changes in hydrophilicity. FTIR-ATR confirmed the unchanged crystallinity of the plasma-treated surfaces. The XPS showed an increase in oxygen in the samples exposed to plasma, along with the detection of phosphate and calcium ions in the functionalized samples. In addition, changes in the topography of each sample related to the different surface treatments were observed using SEM. These surface modifications showed a more favorable cellular response compared with the control. The fibroblast cells showed a progressive increase in viability and proliferation with each subsequent treatment, and at 96 h, they exhibited a similar proliferation pattern. Osteoblast cells showed very close viability in all plasma-treated and functionalized samples and a remarkable preference for proliferation in the surface functionalized with both ions. This indicates that plasma activation supports fibroblast growth, but phosphate–calcium functionalization promotes a more consistent osteogenic response. Regarding cell adhesion, both cell lines showed a more extended morphology in all samples compared with the Control. For this sample, mechanically polished, cell viability and proliferation were observed. However, the cellular response was slower, which could be a disadvantage in the healing process and tissue regeneration.
In conclusion, this study demonstrates that surface modification of inert PEEK enhances cellular responses related to viability, proliferation, adhesion, and spreading. Therefore, it may represent a promising strategy to improve the biological performance of polymeric implants, for instance, by promoting strong tissue bonding and integration. In this context, functionalization with phosphate and calcium ions emerges as a simple and cost-effective option.

Author Contributions

L.V.T.-L.: Conceptualization, methodology, validation, formal analysis, investigation, writing—original draft, visualization. J.S.C.-C.: Conceptualization, validation, formal analysis, resources, writing—review and editing, supervision, funding acquisition. A.L.-V.: Methodology, resources, data curation, project administration, writing—review and editing, visualization. C.A.M.-P.: Formal analysis, resources, supervision, writing—review and editing, visualization. S.Y.R.-L.: Methodology, resources, validation, investigation, writing—review and editing, visualization. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data is contained within the article.

Acknowledgments

The authors thank the Bioscience Department at UTEP and CIMAV for providing access to their facilities and for the technical support received.

Conflicts of Interest

The authors declare no conflicts of interest.

Abbreviations

The following abbreviations are used in this manuscript:
PEEKPoly-ether-ether-ketone
SEMScanning electron microscopy
XPSX-ray photoelectron spectroscopy
FTIR-ATRFourier-transform infrared spectroscopy with attenuated total reflectance
MTT3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide
DMEM-F12Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12
CAMKIICalmodulin-dependent kinase II
NFATNuclear factor of activated T cells
CREBcAMP response element-binding protein
ERK ½Extracellular signal-regulated kinase 1 and 2
PI3KPhosphoinositide 3-kinase
AKTProtein Kinase B (PKB)
BMPBone morphogenic protein
IGF-1Insulin-like Growth factor I
RANKLReceptor activator of nuclear factor kB ligand
OPGOsteoprotegerin
WntWingless/integrated signaling pathway

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Coatings 15 01359 g001
Figure 1. Water contact angle measurements on sample surfaces and their corresponding droplets. Values represent the mean ± standard error (n = 3). **** indicates p < 0.0001 vs. control.
Figure 1. Water contact angle measurements on sample surfaces and their corresponding droplets. Values represent the mean ± standard error (n = 3). **** indicates p < 0.0001 vs. control.
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Figure 2. FTIR-ATR spectra for the control and PL-S samples. Crystallinity [%] = (CI − 0.728)/1.549 × 100.
Figure 2. FTIR-ATR spectra for the control and PL-S samples. Crystallinity [%] = (CI − 0.728)/1.549 × 100.
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Figure 3. SEM images indicating the surface PEEK topography for (a) control, (b) PL-S, (c) Ca-S, and (d) PCa-S samples. Scale bar 1 µm.
Figure 3. SEM images indicating the surface PEEK topography for (a) control, (b) PL-S, (c) Ca-S, and (d) PCa-S samples. Scale bar 1 µm.
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Figure 4. XPS chemical analysis: (a) survey spectrum and peak representation for (b) Ca-S sample Ca2p, (c) PCa-S sample Ca2p, and P2p.
Figure 4. XPS chemical analysis: (a) survey spectrum and peak representation for (b) Ca-S sample Ca2p, (c) PCa-S sample Ca2p, and P2p.
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Figure 5. Cell viability determined by the MTT assay at 24 h for (a) fibroblast and (b) osteoblast cells. The bars denote the standard error. Statistical significance: ** p < 0.01; **** p < 0.0001 vs. control.
Figure 5. Cell viability determined by the MTT assay at 24 h for (a) fibroblast and (b) osteoblast cells. The bars denote the standard error. Statistical significance: ** p < 0.01; **** p < 0.0001 vs. control.
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Figure 6. (a) Fibroblast NIH/3T3 and (b) osteoblast HOS cell proliferation analyzed at 24 and 96 h, (n = 2; 20 fields analyzed per sample). Statistical significance: ns (not significant); ** p < 0.01; **** p < 0.0001 vs. control.
Figure 6. (a) Fibroblast NIH/3T3 and (b) osteoblast HOS cell proliferation analyzed at 24 and 96 h, (n = 2; 20 fields analyzed per sample). Statistical significance: ns (not significant); ** p < 0.01; **** p < 0.0001 vs. control.
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Figure 7. Cell adhesion and spreading after 24 h. Nuclear and cytoskeletal structures were stained with Hoechst and Phalloidin, respectively. Bars are 20 µm.
Figure 7. Cell adhesion and spreading after 24 h. Nuclear and cytoskeletal structures were stained with Hoechst and Phalloidin, respectively. Bars are 20 µm.
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Table 1. Sample labeling.
Table 1. Sample labeling.
LabelNameSurface Treatment
ControlControl sampleNo surface treatment
PL-SPlasma samplePlasma-activated surface
Ca-SCalcium sampleCalcium functionalized surface
PCa-SPhosphate and calcium samplePhosphate and calcium functionalized surface
Table 2. Atomic percentage composition.
Table 2. Atomic percentage composition.
SampleC1sO1sP2pCa2pCa/P
Control79.47 ± 0.1120.54 ± 0.11
PL-S73.27 ± 3.6226.73 ± 3.62
Ca-S72.69 ± 4.6924.92 ± 3.63 2.38 ± 1.12
PCa-S50.5 ± 3.0334.83 ± 1.526.47 ± 0.568.20 ± 1.071.26
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MDPI and ACS Style

Tapia-Lopez, L.V.; Luna-Velasco, A.; Martínez-Pérez, C.A.; Reyes-López, S.Y.; Castro-Carmona, J.S. Enhancing PEEK Surface Bioactivity Through Phosphate and Calcium Ion Functionalization. Coatings 2025, 15, 1359. https://doi.org/10.3390/coatings15111359

AMA Style

Tapia-Lopez LV, Luna-Velasco A, Martínez-Pérez CA, Reyes-López SY, Castro-Carmona JS. Enhancing PEEK Surface Bioactivity Through Phosphate and Calcium Ion Functionalization. Coatings. 2025; 15(11):1359. https://doi.org/10.3390/coatings15111359

Chicago/Turabian Style

Tapia-Lopez, Lillian V., Antonia Luna-Velasco, Carlos A. Martínez-Pérez, Simón Yobanny Reyes-López, and Javier S. Castro-Carmona. 2025. "Enhancing PEEK Surface Bioactivity Through Phosphate and Calcium Ion Functionalization" Coatings 15, no. 11: 1359. https://doi.org/10.3390/coatings15111359

APA Style

Tapia-Lopez, L. V., Luna-Velasco, A., Martínez-Pérez, C. A., Reyes-López, S. Y., & Castro-Carmona, J. S. (2025). Enhancing PEEK Surface Bioactivity Through Phosphate and Calcium Ion Functionalization. Coatings, 15(11), 1359. https://doi.org/10.3390/coatings15111359

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