1.

Introduction

One of the clinical research goals during the last decade has been the clarification of staphylococcal pathogenesis mechanisms as they relate to implanted biomaterials for joint prostheses, catheters, heart valves, and pacemakers.¹ The pathogen which represents a common source of infections on indwelling medical devices is Staphylococcus epidermidis, a gram-positive bacterium that permanently resides on human skin and in mucosa membranes. S. epidermidis is capable of adhering to synthetic biomaterial surfaces and forming a stable biofilm leading to persistent infections.^2,3 During biofilm formation, bacterial cells secrete extracellular polymeric substances that include lipids, proteins, polysaccharides, and others; these protect the bacterial colony from external attack.^4,5 Accordingly, treating biofilm-related infections is extremely difficult. Actions of the immune system can be ineffective and eradication of the pathogen often requires extreme doses of antibiotics. Biofilm-encased bacteria are up to a 1000 times more resistant to bactericidal drugs than their planktonic predecessors.⁶ In addition, for patients with compromised immune systems, the release of bacteria from the biofilm often triggers remote secondary site infections. One possible way of preventing periprosthetic infections is to design biomaterials with physical or chemical barriers that are capable of inherently resisting bacterial surface adhesion.^7,8 One candidate material with such properties is ${\mathrm{Si}}_3{\mathrm{N}}_4$, a nonoxide ceramic whose surface can be chemically and topographically altered.⁹ Previous tests conducted by Bock et al.⁹ verified the antibacterial behavior of this bioceramic. They compared the bacteriostatic characteristics of different biomaterials used in medical devices against S. epidermidis and Escherichia coli (E. coli). Their results showed a pronounced reduction in bacteria adhesion and live bacteria with increased exposure to ${\mathrm{Si}}_3{\mathrm{N}}_4$.

Raman spectroscopy on living bacteria has been employed to follow its metabolism and to understand the chemical interactions between bacterial cells and biomaterials.^10–12 This in situ spectroscopic technique is useful in identifying and observing specific time-dependent biochemical changes in the composition and activity of prokaryotic cells. Neugebauer et al.¹⁰ succeeded in characterizing S. epidermidis with respect to its metabolic functions using different spectroscopic techniques; Samek et al.¹¹ analyzed the formation of S. epidermidis biofilm by labeling characteristic Raman bands in its spectrum. The metabolism of gram-negative Porphyromonas gingivalis was also clarified in a recent paper before and after its inoculation onto various ${\mathrm{Si}}_3{\mathrm{N}}_4$ substrates.¹²

In this study, the in situ metabolic activity, cellular growth, and lysis of S. epidermidis were examined at different time points under the chemical and physical influences of various ${\mathrm{Si}}_3{\mathrm{N}}_4$ substrates. Comparative data were also acquired on two other popular biomaterials: polyetheretherketone (PEEK) and a titanium alloy (Ti). In situ Raman spectroscopy provided physicochemical insights into the interaction of S. epidermidis colonies with these various biomaterials, from which a mechanistic understanding of ${\mathrm{Si}}_3{\mathrm{N}}_4$’s bacteriostatic behavior was proposed.

2.

Materials and Methods

3.

Results

3.1.

Topography of Substrate Surfaces

Laser microscopy images and related three-dimensional (3-D) plots for different substrates are shown in Fig. 1. Untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$ [Fig. 1(a)] exhibited a peculiar topology with the presence of protruding acicular grains interspersed into an oxide intergranular matrix. The ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$ surface [Fig. 1(b)] presented a similar acicular grain structure but also displayed increased areal fractions of the intergranular phase that was expressed from the bulk during the thermal treatment.¹⁵ Figures 1(c) and 1(d) show the surfaces of the Ti-alloy and PEEK, respectively.

Fig. 1

Laser micrographs (left side) and relative topography plots (right side) of (a) untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$, (b) ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$, (c) Ti-alloy, and (d) PEEK. Images are obtained before bacterial treatment at $150\times$ magnification.

The surface topology of the Ti-alloy was regular and homogeneous without the presence of secondary phases; the surface of PEEK appeared to be the smoothest of the tested materials. The average roughness, Ra, for these samples ($n=4$ each) is compared in Fig. 2 together with a statistical analysis of observed differences. The ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$ presented the highest roughness ($0.40\pm 0.04\mu \mathrm{m}$) followed by the untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$ ($0.32\pm 0.02\mu \mathrm{m}$). The PEEK surface possessed the lowest roughness ($0.26\pm 0.03\mu \mathrm{m}$), whereas the topology of the Ti-alloy ($0.29\pm 0.04\mu \mathrm{m}$) was not significantly different from untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$.

Fig. 2

Roughness parameter, Ra, of substrate samples; ${\mathrm{N}}_2$-annealed samples were smoother than other samples (n.s. = non significant; $p<0.05$), whereas untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$ did not show a significant statistical difference when compared with the Ti-alloy ($p>0.05$).

3.2.

Microbial Viability Assays and Fluorescence Microscopy

As shown in Figs. 3(a) and 3(b), bacterial cell viability was verified at 12 and 48 h, respectively. After 12 h of exposure, the ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$ showed the highest OD value ($3.12\pm 0.044$) indicating the largest number of living bacteria among the studied materials. The Ti-alloy had the lowest OD value ($1.38\pm 0.02$), whereas the PEEK ($2.14\pm 0.06$) and untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$ ($2.03\pm 0.08$) had intermediate OD values. OD values between the tested materials were significantly different ($p<0.05$) with the exception of the PEEK and untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$. However, a noteworthy variation in bacterial growth occurred at the 48-h time-point. The highest OD value belonged to PEEK ($0.87\pm 0.09$), whereas the ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$ displayed the lowest OD value ($0.59\pm 0.02$). Again, a comparison between untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$ ($0.73\pm 0.02$) and the Ti-alloy ($0.69\pm 0.04$) resulted in a minimal statistical difference. After 48 h, all samples were statistically different ($p<0.05$).

Fig. 3

WST assay diagram obtained after (a) 12 h and (b) 48 h bacterial treatment with S. epidermidis. After 48 h, all samples showed statistical differences among each other ($p>0.05$).

Note that the OD values measured for all samples were lower at 48 h than at 12 h. This was likely a consequence of the two sets of experiments being performed on independent samples, rather than representing time-series events.

At 24 h of exposure, fluorescence micrographs were obtained for all samples after first staining them with PI, CFDA, and DAPI markers, which colored dead bacteria (red), live bacteria (green), and nuclei (blue), respectively. Results are shown in Fig. 4 with a constant magnification of $20\times$ for all samples. For the silicon nitride substrates, a relatively high amount of bacteria was detected [cf. DAPI blue-stained micrographs in Figs. 4(a) and 4(b)] for both the untreated and ${\mathrm{N}}_2$-annealed samples, respectively. However, PI red-stained areas associated with dead bacteria clearly indicated that significant lysis had occurred, and it was more evident on the ${\mathrm{N}}_2$-annealed material. Red stains were also detected on the Ti-alloy [Fig. 4(c)], with its surface presenting a relatively low density of bacterial cells. Micrographs on PEEK [Fig. 4(d)] confirmed no antimicrobial behavior for this material. There were no apparent dead bacteria and a high number of living cells over the entire sample’s surface, indicating significant bacteria proliferation.

Fig. 4

Fluorescence micrograph after PI, CFDA, and DAPI staining of S. epidermidis exposed for 24 h to: (a) untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$, (b) ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$, (c) Ti-alloy, and (d) PEEK. Live and dead cells were labeled with green and red stains, respectively, whereas nuclei displayed in blue color. Images were obtained at a $20\times$ magnification.

3.3.

Raman Spectroscopy Results

3.3.1.

Labeling Raman emissions from control samples

Figure 5 shows average Raman spectra collected on living S. epidermidis bacteria cultured on a silica glass control substrate. Three spectra are shown as a function of exposure time (12, 24, and 48 h; as labeled in the inset) in the spectral region between 300 and $3500{\mathrm{cm}}^{-1}$. All the observed bands belonged to molecular species within S. epidermidis. The spectra were divided into four main regions, labeled zones I to IV, and individual bands were assigned according to their physical origin and vibrational modes in the cellular compounds (cf. Fig. 5 and Tables 1Table 2–3).

Fig. 5

Raman spectra of S. epidermidis ATCC 14990 in the 350 to $3500{\mathrm{cm}}^{-1}$ range collected after different exposure times to a silica glass surface used as a control sample for bacteria metabolic evolution. Spectra were divided into four distinct zones (labeled zones I to IV), while the main band frequencies were labeled according to Tables 1–3.

Table 1

Band labeling, assignment, and related references in spectral zone I.

Band	Assignment	Frequency (cm−1)	Reference
1	Skeletal vibrations in tyrosine	638	17, 18
2	Ring vibrations in poly-deoxyadenosine	650	14, 15
3	Ring breathing in guanine (DNA)	670	14, 15
4	Phosphatidylserine	720	14
5	Ring breathing adenine (DNA)	735	15
6	Ring breathing in tryptophan (DNA)	760	17, 18
7	Phosphodiester stretching	782	15
8	Out-of-plane ring breathing in DNA	820	17, 18
9	COC 1,4 glycosidic link in carbohydrates	840	16
10	Ring breathing in trypsin (DNA)	850	14

Table 2

Band labeling, assignment, and related references in spectral zone II.

Band	Assignment	Frequency (cm−1)	Reference
11	$\mathrm{C}─\mathrm{O}─\mathrm{C}$ glycosidic linkage of carbohydrates	937	19
12	$\mathrm{C}─\mathrm{C}$ stretching in $\beta$-sheet	976	17, 18
13	Ring breathing in phenylalanine	1004	17, 18
14	$\mathrm{C}─\mathrm{C}$gauche stretching in membrane lipids	1092	19
15	$\mathrm{C}─\mathrm{C}$trans stretching in membrane lipids	1125	19
16	$\mathrm{C}─\mathrm{N}$ stretching amide III in $\beta$-sheet	1253	20
17	Twisting of saturated ${\mathrm{CH}}_2$ in membrane lipids	1313	21
18	Symmetric stretching of $\mathrm{O}─\mathrm{C}─\mathrm{O}$ in ${\mathrm{COO}}^{-}$	1400	22
19	Bending of saturated ${\mathrm{CH}}_2$ in membrane lipids	1442	21

Table 3

Band labeling, assignment, and related references in spectral zones III and IV.

Band	Assignment	Frequency (cm−1)	Reference
20	$\mathrm{C}═\mathrm{O}$ stretching amide I ($\alpha$-helix)	1600	15
21	$\mathrm{C}═\mathrm{O}$ stretching amide I (degraded structure)	1630	27
22	$\mathrm{C}═\mathrm{O}$ stretching amide I ($\beta$-sheet)	1680	23

In zone I, between 600 and $900{\mathrm{cm}}^{-1}$, a set of bands related to DNA and RNA dominated the Raman spectrum. In particular, the band at $720{\mathrm{cm}}^{-1}$ was attributed to adenine containing species, whereas bands at 735 and $782{\mathrm{cm}}^{-1}$ were related to a mode of adenine ring breathing in DNA and DNA phosphodiester stretching, respectively.^17,18 Additional bands in this region were located at 820 and $840{\mathrm{cm}}^{-1}$, the former corresponding to out-of-plane ring breathing modes of tyrosine and the latter to the $\mathrm{C}─\mathrm{O}─\mathrm{C}$ stretching vibration of 1,4 glycosidic link in carbohydrates.^19–21

Zone II at 900 to $1500{\mathrm{cm}}^{-1}$ was characterized by a series of emission bands from amides, lipids, and carbohydrates. Two main bands in this region, at 1092 and $1125{\mathrm{cm}}^{-1}$, are related to $\mathrm{C}─\mathrm{O}─\mathrm{C}$ antisymmetric stretching in aliphatic esters and glycosidic link of carbohydrates, and proteins, and lipids, respectively.²² Additional bands at 1253 and $1313{\mathrm{cm}}^{-1}$ are related to $\mathrm{C}─\mathrm{N}$ stretching in amide III and twisting of saturated ${\mathrm{CH}}_2$ moieties in membrane lipids, respectively.^23,24 Another prominent band at $1442{\mathrm{cm}}^{-1}$ corresponds to bending of saturated ${\mathrm{CH}}_2$ in polysaccharides and membrane lipids.^24,25

Zone III between 1500 and $1850{\mathrm{cm}}^{-1}$ contained one $\alpha$-helix-related band ($\mathrm{C}═\mathrm{O}$ stretch) at $1600{\mathrm{cm}}^{-1}$,¹⁸ but it was dominated by the $\beta$-sheet-related $\mathrm{C}═\mathrm{O}$ stretching emission of amide I at $1680{\mathrm{cm}}^{-1}$.²⁶

Zone IV between 2800 and $3100{\mathrm{cm}}^{-1}$ contained three main bands at $2880{\mathrm{cm}}^{-1}$ (symmetric stretching of ${\mathrm{CH}}_3$ in lipids and fatty acids²⁷), $2934{\mathrm{cm}}^{-1}$ (symmetric and antisymmetric C-H stretching of ${\mathrm{CH}}_2$ and ${\mathrm{CH}}_3$ in lipids, proteins, and carbohydrates²⁸), and $2970{\mathrm{cm}}^{-1}$ (assigned to ${\mathrm{CH}}_3$ symmetric stretching in lipids and fatty acids²⁹).

Looking at the time-lapse spectra collected on the control substrate revealed a slight intensity decrease of the triplet at 2881, 2934, and $2970{\mathrm{cm}}^{-1}$ and of the amide I band at $1670{\mathrm{cm}}^{-1}$ at 24-h exposure as compared with 12 h. However, the spectrum at 48 h did not show significant variations as compared with 24 h or any significant decreases in band intensities. Rather, DNA bands of phosphodioxy groups and adenine at 1092 and $729{\mathrm{cm}}^{-1}$ experienced higher intensities after 48 h than after 24 h. Another noted difference was the amide I band at $1680{\mathrm{cm}}^{-1}$. It was slightly more intense at 48 h compared with 24 h. A similar trend was observed for protein and carbohydrate bands at 836 and $858{\mathrm{cm}}^{-1}$; these were completely absent after 24 h but were clearly visible after 48 h. The evolution of these bands as a function of time on the silica substrate was utilized as a standard for the metabolic activity of S. epidermidis. It was used to establish meaningful criteria for judging changes in the bacteria’s metabolism upon its exposure to the test samples. It should be noted that bands related to proteins and carbohydrates actually change with the formation of a biofilm, whereas DNA-related bands are barometers of bacterial cell proliferation and well-being.

3.3.2.

Raman emissions from S. epidermidis on different substrates

Normalized Raman spectra of the bacteria as a function of exposure time on the different substrates in zone I are shown in Figs. 6(a)–6(c) for 12, 24, and 48 h, respectively. As shown in Table 1,^{17,18,19–21} 10 distinct bands were found and a number of features became visible in their time dependencies. In comparison with the control sample, the emission at $720{\mathrm{cm}}^{-1}$ (labeled as band 4 and related to phosphatidylserine) tended to disappear for both untreated and ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$. In contrast, this band underwent a decrease in intensity at >24 h for bacteria exposed to the Ti-alloy. In the main triplet of bands 5 to 7, only the phosphodiester band 7 completely disappeared with elapsed time for both ${\mathrm{Si}}_3{\mathrm{N}}_4$ substrates, whereas the DNA-related band 6 increased in relative intensity for both ${\mathrm{Si}}_3{\mathrm{N}}_4$ samples. Band 7 persisted up to 24 h with a relatively strong intensity for the Ti-alloy substrate. Band 8 tended to disappear upon exposure of the bacteria to untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$, whereas it showed an increase at 24 h and then a reduction at 48 h in the case of ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$. The trend found for band 8 in bacteria exposed to the Ti-alloy substrate was similar to that observed for ${\mathrm{N}}_2$-annealed substrate. Band 10 (ring breathing in trypsin) was quite strong for bacteria exposed to both types of ${\mathrm{Si}}_3{\mathrm{N}}_4$ substrates, whereas it almost disappeared after 48 h on the Ti-alloy.

Fig. 6

Normalized and deconvoluted Raman spectra in spectral zone I from S. epidermidis exposed to different substrates (cf. labels in the inset to each column) for (a) 12 h, (b) 24 h, and (c) 48 h.

Spectral emissions in zone II [Figs. 7(a)–7(c) for 12, 24, and 48 h exposures, respectively] are shown in Table 2.^17–21

Fig. 7

Normalized and deconvoluted Raman spectra in spectral zones II and III from S. epidermidis exposed to different substrates (cf. labels in the inset to each column) for (a) 12 h, (b) 24 h, and (c) 48 h.

They included nine bands labeled as bands 11 to 19. In this zone, a clear difference was noted in the trend of the phenylalanine band 13 at $\sim 1004{\mathrm{cm}}^{-1}$ for the glass control, the two ${\mathrm{Si}}_3{\mathrm{N}}_4$ materials, and Ti-alloy. Although in the case of glass control, this band maintained a moderate relative intensity with elapsing exposure in comparison with neighboring bands, its intensity was quite pronounced for both of the ${\mathrm{Si}}_3{\mathrm{N}}_4$ substrates and almost null for the Ti-alloy. Bands 17 to 19, which are contributed by membrane lipids, showed an abnormal increase in their relative intensities for S. epidermidis at intermediate exposures to the ${\mathrm{N}}_2$-annealed substrate. A similar trend, although less pronounced, was observed for untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$ and Ti-alloy. The two bands sensitive to lipid conformation, namely bands 14 and 15, showed a clear inverse trend in their relative intensity upon exposure to untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$ and the Ti-alloy as compared with the glass control. This change consisted of a significant increase in the latter band, which can be interpreted as a lipid population with a trans versus a gauche configuration.

Amide I dominated zone III [1500 to $1750{\mathrm{cm}}^{-1}$; Figs. 7(a)–7(c); Table ^18,25,29,30] with its main feature being band 21 ($\mathrm{C}═\mathrm{O}$ stretching emission at $1680{\mathrm{cm}}^{-1}$). This band followed the same trend observed for the lipid-related bands 17 to 19 in zone II, namely a significant enhancement at intermediate exposure times for both ${\mathrm{Si}}_3{\mathrm{N}}_4$-based substrates, but it was more pronounced for the ${\mathrm{N}}_2$-annealed material. An enhancement in the relative intensity of band 21 was also observed for the Ti-alloy as compared with glass control. However, no significant differences were noted as a function of exposure time up to 48 h.

4.

Discussion

Mechanisms of bacterial adhesion onto and interaction with biomaterial surfaces are complex and depend on many factors, such as morphological and chemical features of the surfaces. Surface topography plays a fundamental role in bacteria/substrate interaction affecting both adhesion and biofilm formation. In general, rough surfaces promote adhesion and colonization.³¹ For example, Yoda et al.³² tested the bacteriostatic behavior of different substrates with controlled surface roughness with respect to S. epidermidis. They showed that adhesion and proliferation were greatly favored on rougher materials. However, nanorough surfaces have an opposite effect. Inspired by nature, the nanostructured surfaces of cicada wings³³ have led to the development of engineered antimicrobial features on various biomaterials including titanium, polyurethane, and PEEK.^34–37 Additionally, as demonstrated by two independent studies, the nanofeatures of untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$ proved to be more resistant to bacterial adhesion and biofilm formation than chemically identical substrates that were either machined or polished.^38,39

In this study, the ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$ presented the highest Ra value among the tested materials (cf. Fig. 2). It also showed the highest amount of living cells after 12 h according to microbial viability tests [cf. Fig. 3(a)]. Conversely, the PEEK substrate had the smoothest surface among the tested substrates, but the amount of living bacteria on its surface after 12 h was comparable with the untreated ${\mathrm{Si}}_3{\mathrm{N}}_4$ and Ti-alloy despite the fact that these latter materials experienced substantially higher Ra values. The situation after 48 h was found completely reversed with the ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$ possessing the lowest quantity and the PEEK having the highest amount of living bacteria among the tested samples. These data suggest that the bacteriostatic behavior of the tested surfaces cannot simply be interpreted using surface roughness data alone. Surface chemistry needs to be taken into consideration as well. The chemical reactions at the surface of ${\mathrm{Si}}_3{\mathrm{N}}_4$ were discussed in a previous study.¹² They suggested the existence of a diffusion-limited thermodynamic driving force to convert silicon nitride into silicic acid ($\mathrm{Si}{(\mathrm{OH})}_4$) and ammonia (${\mathrm{NH}}_3$). Release of ${\mathrm{NH}}_3$ from the ceramic surface was experimentally demonstrated by a significant increase in local pH (i.e., from $\sim 5.5$ to $\sim 8.5$).¹² As indicated by prior research, highly alkaline environments may delay or prevent bacterial adhesion and biofilm formation. For instance, Hamadi et al.⁴⁰ investigated the adhesion of Staphylococcus aureus to glass at different pH values and observed that bacteria strongly adhered in the pH range 4 to 6, whereas adhesion was weak under both highly acidic ($\mathrm{pH}\approx 2$ to 3) and alkaline conditions ($\mathrm{pH}>7$).

Raman spectroscopy provided further insight into the bacteria’s metabolism by showing the time-dependent variations of its molecular structure after contact with the different substrates. A significant reduction in the intensity of its DNA-related bands in spectral zone I (e.g., bands 2, 3, 7, and 8) after 48 h of exposure to both types of ${\mathrm{Si}}_3{\mathrm{N}}_4$ and the Ti-alloy substrates was an important indicator of its metabolic degradation. Escoriza et al.⁴¹ studied the time-lapse Raman response of bacteria during their proliferation and found that bands related to nucleic bases typically showed high intensities at the beginning of bacterial growth, but their intensities slowly decreased with time. In the current experiments, a similar initial trend coupled with an abrupt decrease at 48 h was observed. This sudden decrease strongly suggests that both ${\mathrm{Si}}_3{\mathrm{N}}_4$ materials and the Ti-alloy possess inherent antibacterial mechanisms.

Alterations of phospholipid-related and protein-related bands were also observed, which indicated major irreversible modifications to the bacteria’s membrane and protein structures. The spectral bands at $1125{\mathrm{cm}}^{-1}$ (band 15) and $1442{\mathrm{cm}}^{-1}$ (band 19) monitor the polysaccharides for intercellular adhesion and biofilm formation.^42,43 These bands initially increased but then conspicuously disappeared after exposure to ${\mathrm{N}}_2$-annealed ${\mathrm{Si}}_3{\mathrm{N}}_4$ for 48 h. This trend can be interpreted as disruption of the biofilm after its initial formation at intermediate times. An important marker for proteins is band 13 at $1004{\mathrm{cm}}^{-1}$ related to vibrational modes in phenylalanine. However, this band tended to disappear only for bacteria exposed to the Ti-alloy. The strong band 17 (at $1313{\mathrm{cm}}^{-1}$) reflects twisting vibrations of saturated ${\mathrm{CH}}_2$ bonds in membrane lipids. When comparing the relative intensity of this band to that of phenylalanine (band 13), a significant enhancement was noticed after 24 h of exposure to both types of ${\mathrm{Si}}_3{\mathrm{N}}_4$ surfaces, followed by a clear intensity reduction after 48 h. The intensity enhancement of bands 17 was interpreted as acidification of the membrane lipids.²⁴ However, bacteria have an ability to control the biophysical properties of the phospholipids in their membranes in the presence of an external physiological stress⁴⁴ and this was observed via Raman spectroscopic monitoring in the current experiments. Following the work of Nostro et al.,⁴⁵ the bacterial behavior observed on the ${\mathrm{Si}}_3{\mathrm{N}}_4$ substrates was interpreted as a reaction to the increase in local pH. This reaction reduced the bacteria’s initial adhesion capability and subsequently led to the impairment of biofilm maturation. An alkaline pH directly influences bacterial attachment because it affects the degree of D-alanylation of lipoteichoic acid and wall teichoic acid, which are the most abundant polyanions in cytoplasmic membrane lipids and are linked to peptidoglycan. Another important aspect of the change in pH is its impact on enzymes related to autolysis of prokaryotic cells. A pH change favors activation of autolysin leading to peptidoglycan hydrolases, which strongly alters peptidoglycan, turn over. When hydrolase is activated, peptidoglycans are degraded and the osmotic balance of the bacterial cell is lost with the cell turning from a respiring to a depolarized status. Ammonium moieties released from the ${\mathrm{Si}}_3{\mathrm{N}}_4$ surface¹² then become free to flow into the cytoplasmic environment because the structural integrity of the cell wall is compromised. This process ultimately results in cell lysis.⁴⁶ Figure 8 shows a schematic model of S. epidermidis exposed to a silicon nitride surface with a time-dependent increase in local pH. A process of uncontrolled autolysis is envisaged, which modifies the peptidoglycan wall structure and increases the susceptibility of the membranes to molecular moieties liberated by the substrate, thereby destabilizing the cell.

Fig. 8

(a) Schematic model of S. epidermidis bacterial membrane in osmotic equilibrium and (b) in depolarized status after being affected by polyglycan hydrolase.

5.

Conclusion

To clarify the origin of ${\mathrm{Si}}_3{\mathrm{N}}_4$’s bacteriostatic behavior, changes within gram-positive S. epidermidis were studied after its exposure to substrates consisting of two ${\mathrm{Si}}_3{\mathrm{N}}_4$ variants, a Ti-alloy and a glass control for time-points up to 48 h. Previously reported results were confirmed using microbial viability assays and fluorescence microscope analyses. Then, in situ Raman spectroscopy was employed to monitor the metabolic variations of the bacteria at different exposure times. These variations were interpreted based on the vibrational responses of their biological DNA. The Raman spectra showed a decrease in the band intensities related to biofilm formation and DNA after 48 h suggesting the initiation of lysis. The Raman data agreed with the results derived from the biological tests. Based on these observations and in comparison with previously published literature, the metabolic changes in S. epidermidis were attributed to a pH drift toward alkaline values on the ${\mathrm{Si}}_3{\mathrm{N}}_4$ surfaces. This drift modified the behavior of enzymes and cellular compounds, limiting its adhesion and increasing the permeability of the cell walls. In conclusion, this study demonstrated that the surface chemistry of ${\mathrm{Si}}_3{\mathrm{N}}_4$ may be useful in inhibiting periprosthetic infections by hindering biofilm formation and bacterial proliferation.