Effects of multiple implantations of titanium healing abutments: Surface characteristics and microbial colonization

Abstract

Objective.

Very few studies have investigated dental implant components involved in the early stage of healing, especially the implant healing abutment (IHA), despite its vital role in soft tissue contouring and shaping after implant placement. Although these components are labelled by the manufacturer for “single-use only,” it is a common clinical practice to clean, sterilize, and reuse them.

Methods.

In the present study, IHAs after single and multiple implantations were retrieved as per standard procedures, and biological material isolated from the surface was subjected to 16S rRNA sequence analysis. The microbiome analysis was followed by cleaning and sterilization in order to replicate clinical sterilization techniques. Following sterilization, retrievals were subjected to surface characterization with optical and scanning electron microscopy to investigate surface features, and electrochemical testing was performed to evaluate corrosion behavior.

Results.

The microbiota was comprised of early colonizers including Streptococcus species and secondary anaerobic colonizers such as Fusobacterium, Capnocytophaga, and Prevotella species. The surface analysis revealed that irrespective of the cleaning and sterilization techniques, the pristine, homogeneous surface of the new, unused IHAs could not be restored. Both single and multiple-use IHAs had severe surface changes including discoloration, major abrasions, biological contamination, and the IHA retrievals exhibited higher corrosion rate as compared to control specimens.

Significance.

Reusing IHAs multiple times may not be a prudent practice as the microbial colonization and surface changes caused by using this component multiple times may affect the performance of IHAs in soft tissue healing.

1. Introduction

Bacterial adhesion plays a significant role on dental implant performance as it can impact both hard and soft tissues, with colonization occurring within minutes following implant placement [1–3]. It has been suggested that a successful dental implant outcome can be predicted based on a “race for the surface” between oral bacteria and soft tissue cells [4]. In this regard, other components of the dental implant system are exposed to these conditions where the competition between bacteria and host cells can impact tissue healing and later implant integration. One component of the dental implant system, which is often overlooked due to its temporary placement, is the healing abutment. The implant healing abutment (IHA) is an important component as it remains exposed to the oral environment during the early healing phase. Thus, the IHA is exposed to a unique combination of conditions, with one-part supragingival and exposed to the oral cavity and the other part subgingival and in contact with soft tissue. In addition, this component is essential for soft tissue conditioning as it provides a scaffold for tissue growth [5].

IHAs typically have smooth machined surfaces and are composed of commercially pure titanium (cpTi) due to its biocompatibility, corrosion resistance, and excellent mechanical properties [6,7]. These devices often undergo a surface treatment called anodization which provides additional resistance to corrosion and better protects the bulk material when exposed to corrosive substances from diet, human saliva, and oral biofilms. The different types of IHA commercially available are chosen based on implantation procedures and specific protocols used by clinicians. The IHAs used in this study included both bone-level and tissue-level implants.

Placement of an implant body and the supporting components like the IHA subject the jaw bone and soft tissue to surgical trauma. This results in a cascade of immune responses both in the bone and surrounding soft tissues. The IHA is known to help in the formation of a long-standing biological barrier, thus allowing soft tissue healing after the placement of a dental implant body [8]. Hence, the successful interface of IHA with soft and hard tissues during the early healing period is necessary to reduce complications that can later affect implant integration [9].

In the literature, early implant complications are reported to occur due to synergistic factors like overloading or surgical error, micromotion, and bacterial contamination resulting in postoperative infection of soft and hard tissues [9]. These complications can be initiated by surface adhesion of early bacterial colonizers which upon the formation of a biofilm may prevent sealing of epithelial tissues on the implant surface. The colonization of these pathogens leads to the formation of mature plaque and is considered to be the causative agent of later events such as peri-implantitis development. In particular, bacterial infiltration is a primary reason for both early and late stage complications [10,11]. At the implant body-IHA connection, bacterial leakage can contribute to corrosion and structural damage [12].

The adhesion of bacterial biofilms at early stages post-implantation on implant components is of concern because it can alter the electrochemical conditions of the surface, especially in crevice areas around dental implant components. These peri-implant crevices are found to have some of the periodontopathogens in high abundance [13,14]. A reduction in pH due to the by-products of bacterial metabolism and other inflammatory processes (e.g., peri-implantitis, perimucositis) can create an acidic micro-environment that is ideal for Ti oxidation [15]. Specifically, the acidity of the micro-environment within crevice areas accelerates the dissolution of the passive oxide layer thereby causing localized destruction and crevice corrosion [16,17]. Moreover, pathogens residing within crevices due to implant modularity, such as at implant body-IHA surface, can produce corrosion products, which can potentiate the host inflammatory response and potentially trigger early complications [12,16]. Adherent bacteria can also generate crevice regions between the developing biofilm and the implant component surface [14]. Hence, the role of bacteria present in the early healing period and their effects on the surface features of IHAs must be further investigated.

The current clinical practice is to clean, sterilize and re-use IHAs. The cleaning and sterilization procedures are not standardized but usually include steps such as mechanical wiping, ultrasonication, and steam autoclaving [18–21]. However, such cleaning procedures may not be effective for IHAs. For instance, the screw thread could undergo bioburden despite ultrasonication which cannot remove all biological debris accumulated in the notch area. Plaque accumulation within the notch regions could affect the locking of IHA onto the implant body [22]. Furthermore, several studies have indicated that a combination of mechanical and chemical cleansing is ineffective in complete removal of biological debris and biofilm from implants/abutments [19,20,23,24]. Also, multiple cycles could affect the biocompatibility of IHA surfaces and could result in fracture of the temporary components [19,20]. In addition, a retrieval study showed the presence of viable bacteria attached to IHA surfaces post-sterilization [25]. Thus, utilizing used, cleaned IHAs on multiple patients could possibly result in cross-contamination and impaired healing. In order to assess the overall effectiveness of reusing IHAs, factors favoring the successful functioning of this temporary component need to be further investigated.

The goals of this study were to identify bacterial taxa colonizing IHAs during the early healing period and to elucidate the effects of multiple implantations on the surface characteristics of the IHA. The conjoint results of surface characterization and microbiome analysis can aid in better understanding of changes that IHAs undergo during the initial healing phase and the prospective adverse effects, if any, that may influence soft tissue health. Thus, the importance of this component should not be overlooked; in fact, the IHA can serve as a model to understand the microbiota of the early healing period and the competition between host cells and bacteria during this stage on implant components. The current study also provides insight into the unstable conditions of the oral cavity, post placement, for the design of subsequent in vitro studies. Given the rationale for this study, it was hypothesized that IHA usage may have a role in microbial colonization dynamics. The single- and multiple-use IHAs would exhibit higher corrosion susceptibility and surface damage as compared to unused IHAs.

2. Materials and methods

2.1. Implant healing abutment sample preparation

Sixteen titanium IHAs (Straumann LLC, Basel, Switzerland) were obtained through a private dental practice in Dallas, TX. These components were received from human patients at periods of 3–6 months post-dental implant placement for research purposes, following standard procedures as per guidelines of the Helsinki Declaration. Prior to IHA removal, all patients signed a written informed consent to participate in the study. Patient information remained confidential with only the clinicians having access to health records and demographic information. Retrieval procedures, documentation, and IHA characterization were performed with strict guidelines and were approved by the Institutional Review Board at The University of Texas at Dallas (ER IRB #16–65). All the surgeries were performed by one calibrated surgeon to eliminate the variability produced by different surgical approaches. The retrievals were classified based on the number of implantations to which they were subjected. IHAs that were only used once were grouped as “single-use”. IHAs known to have been used more than once were labeled as “multiple-use”. For multiple-use IHAs, no records were maintained regarding the exact number of times the devices were implanted on different patients. All IHAs obtained post-implantation were marked with a letter for identification (A-P). Four unused IHAs were used as controls for surface characterization and electrochemical testing. Information about the IHAs used in this study is detailed in Table 1.

Table 1 –

Description of implant healing abutments (IHAs) studied.

IHA	Use	Patient number	Type	Connectiona	Corrosion score (1–3)
A1	Multiple	1	Bone	RC	3
D1	Multiple	2	Bone	NC	3
E1	Multiple	2	Bone	RC	3
F	Multiple	3	Bone	RC	3
G	Multiple	4	Bone	RC	1
I	Multiple	5	Bone	RC	2
K	Multiple	6	Bone	NC	3
L	Multiple	7	Bone	RC	2
Bb	Single	8	Bone	RC	1
C	Single	9	Bone	RC	Not tested
H	Single	10	Bone	RC	3
J	Single	11	Tissue	WN	2
M	Single	12	Tissue	WN	1
N1	Single	12	Tissue	WN	1
Ob	Single	13	Bone	RN	2
P	Single	14	Bone	NC	1

^aRC = regular CrossFit; NC = narrow CrossFit; WN = wide neck; RN = regular neck.

^bIHAs that were excluded from microbiome analysis.

2.2. Microbiome analysis

2.2.1. Sample preparation and gDNA isolation

Immediately following retrieval, 16 IHAs were subjected to preparation steps for 16S rRNA sequence analysis. First, each IHA was immersed in 1.5 mL of 1X phosphate buffered saline (PBS) solution in a microcentrifuge tube. After 30–60 min, IHAs were subjected to vortexing for 30–60 s to detach adherent bacteria from the surface. The solution was centrifuged for 5 min. at 1200 RPM to concentrate the bacteria. Total genomic DNA (gDNA) was isolated from the vortexed solution using the MO BIO Ultraclean Microbial DNA Isolation Kit as per the manufacturer’s protocol (MoBio, CA, USA).

2.2.2. Polymerase chain reaction (PCR) and 16S rRNA sequencing

Of the 16 IHAs, 14 had detectable gDNA and were utilized for 16S rRNA sequence analysis. Illumina MiSeq library preparation and sequencing were performed at Molecular Research LP DNA Institute (Shallowater, TX). Briefly, the hypervariable V1–V3 region of the 16S rRNA gene was amplified using the 27F and 519R primers, with a barcode corresponding to the forward primer [26]. DNA amplification was done using the HotStarTaq Plus Master Mix Kit (Qiagen, CA, USA) under the following PCR conditions: initial denaturing at 94 °C for 3 min, followed by 33 cycles of denaturing at 94 °C for 30 s, primer annealing at 53 °C for 40 s, elongation at 72 °C for 1 min, followed by a final elongation step at 72 °C for 5 min. Samples originating from the same DNA template were pooled together following PCR amplification. Samples were purified using calibrated Ampure XP beads (Agencourt Biosciences, Beverly, MA) and used to prepare the Illumina DNA library. Illumina MiSeq sequencing was performed per the manufacturer’s guidelines.

2.2.3. Sequencing analysis and taxonomic assignment

Microbiome analysis was done with the QIIME 2 version 2018.11 pipeline [27,28]. Sequences were demultiplexed, and Divisive Amplicon Denoising Algorithm 2 (DADA2) joined the paired-end reads, removed chimeras, and clustered the reads into Amplicon Sequence Variants (ASV) [29]. Taxonomy was assigned to the ASVs using the SILVA high quality ribosomal RNA database (v.132). A heat map was generated using GraphPad Prism (v.8.2) by calculating the relative frequency of each taxa identified on each IHA. To identify taxa enriched in multiple-use or single-use IHAs, the following thresholds were applied: (1) at least 90% of the total reads obtained for the taxon must originate from either multiple-use or single-use IHAs; and (2) the taxon was detected in at least 50% of all multiple-use or single-use IHAs. Taxa for which >90% of reads originated from a single IHA sample were excluded.

2.2.4. Statistical analysis

Diversity analysis was calculated using QIIME 2 (v. 2018.11). Rarefication to improve even sampling was set at 1575 sequences per sample, and all of the IHAs used for micro-biome analysis met this requirement (n = 14). Within sample diversity analysis (α-diversity) was assessed by quantifying the Shannon Diversity Index (H) and Pielou’s evenness index. The Shannon Diversity Index accounted for species richness and abundance, and Pielou’s evenness index identified how evenly distributed the ASVs were. Diversity between samples was assessed by both qualitative and quantitative phylogenetic β-diversity metrics. PCoA plots were generated with an unweighted and a weighted UniFrac [30–32]. The unweighted UniFrac calculated diversity by the presence or absence of ASVs, and the weighted UniFrac accounted for the abundance of the present ASVs. Significance was assessed by the two-tailed t-test in GraphPad Prism (v.8.2).

2.3. Surface analysis

2.3.1. Cleaning and sterilization

After recovery of gDNA for microbiome analysis, IHA retrievals were subjected to cleaning and sterilization to replicate clinical practice. IHAs were ultrasonicated with acetone, deionized (DI) water and 70% ethanol for 15–20 min each and steam autoclaved (20 min exposure, 2 h drying).

2.3.2. Optical microscopy

Optical microscopy (OM) was performed to observe topographical changes on the surface of the IHAs following cleaning and sterilization (Keyence VHX-2000). The surfaces of the samples were imaged at magnifications ranging from 25× to 500×.

2.3.3. Scanning electron microscopy

In order to evaluate the surface morphology of the specimens after single or multiple implantations, scanning electron microscopy (SEM) was performed. Areas of interest included those with evidence of chemical or physical degradation, which resulted in visible corrosion or wear on the surface of the IHA. An EVO LS 15 environmental scanning electron microscope (ZEISS, Oberkochen, Germany) was used in high vacuum mode and an accelerating voltage of 20 kV. Samples were observed at magnifications ranging from 20× to 2000×.

2.3.4. Qualitative scoring

Based on visual observations derived from OM and SEM, IHA surfaces were scored and classified in terms of accumulation of biological debris and visible corrosion. The scoring ranged from 0 to 3, with 0 corresponding to no surface damage and 3 corresponding to the highest amount of surface damage as detailed in Table 2. For biological debris accumulation, spots of debris, blood remnants, calcium deposits and bacterial plaque were identified and scored based on their fre quency. Also, changes in the surface morphology in the form of discoloration, abrasions, and apparent surface roughness were labeled as surface degradation. For corrosion scoring, the relative amount of surface area containing discoloration, delamination and other deformities were scored.

Table 2 –

Qualitative scoring of the IHA surfaces.

Score	Biological debris	Corrosion (% of surface area)
0	None	None
1	Low, sporadic	0–15, low
2	Moderate, numerous	16–30, medium
3	High, continuous	>30, severe

2.4. Electrochemical testing

Electrochemical analysis post-surface characterization of IHAs was performed in order to evaluate the corrosion behavior of the specimens. Electrochemical testing of all specimens was performed using guidelines modified from ASTM F2129–15 [33] with an Interface 1000 Potentiostat (Gamry Instruments Inc., Warminster, PA). Electrochemical tests were done using a standard three-electrode electrochemical cell. The reference electrode was a saturated calomel electrode (SCE), the counter electrode was a graphite electrode, and the IHA specimens acted as the working electrode. The IHA specimens were mounted to alligator clips and insulated with electrical tape and thoroughly sealed with MICCROStop (Tolber Chemical Division, Hope. AR) to isolate the region of interest and to avoid the interference of the clip or other areas of the IHA. All samples were immersed in 5 mL of the electrolyte, 1X phosphate buffered saline (PBS), which was maintained at 37 °C. Per sample, three tests were performed in a sequential order: open circuit potential (OCP), linear polarization resistance tests, and anodic Tafel polarization. The OCP was measured for 1 h, and the final value was recorded as the corrosion potential (Ecorr). Next, each sample was anodically polarized from −10 mV to 10 mV vs. Ecorr at rate of 0.1667 mV/s as per the guidelines of ASTM G59–05 [34]. The slope of the plotted applied potential vs. current measurements were recorded as the polarization resistance (Rp). Lastly, the corrosion current (Icorr) was extrapolated from anodic Tafel plots, using the linear region of the plot obtained during sweeping from 0 to 250 mV vs. Ecorr at a scan rate of 1 mV/s. This value was used to calculate corrosion rate (CR) using equations derived from ASTM G102–89 [35]. The Rp and CR values were normalized with each sample’s exposed surface area. The results were further compared between unused (control) (n = 4), single-use (n = 7), and multiple-use (n = 8) IHAs.

2.4.1. Statistical analysis

Statistical analysis was performed for the electrochemical parameters, namely corrosion potential (Ecorr) polarization resistance (Rp), and corrosion rate (CR), using a one-way analysis of variance (ANOVA) followed by post hoc Tukey tests. Both tests were run using GraphPad Prism 7.0 Software (GraphPad Software Inc., USA) at a significance level (α) of 0.05.

3. Results

3.1. Microbiome analysis

After joining of paired-end reads, quality control, and filtering, a total of 97,473 reads corresponding to variable regions (1–3) of the 16S rRNA gene were obtained across all IHAs. The frequency of reads per sample ranged from 1575 to 18,704 sequences. A total of 35 different taxa were identified after aligning the ASVs to the reference database. Supplemental data 1 shows the distribution of reads for each taxon and IHA as well as the results of the enrichment analysis. Of the total reads, 68% were obtained from multiple-use IHAs and 32% from single-use IHAs (Supplemental data 1). The genera Fusobacterium, Streptococcus, Capnocytophaga, and Prevotella were the most abundant taxa across all IHAs and accounted for 66% of the total reads obtained (Fig. 1).

Fig. 1 –

Heatmap of the most abundant taxa across multiple-use (left) and single-use (right) IHAs. Abundance was calculated by dividing the number of reads for each taxon found on an IHA by the total number of reads for that IHA. Abundance is color-coded from low (blue) to high (red). (O), order; (C), class (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article).

Two thresholds and one exclusion criterion were applied to identify taxa enriched in either multiple- or single-use IHAs. For thresholding, at least 90% of the total reads obtained for a taxon had to originate from either multiple-use or single-use IHAs, and that taxon had to be detected in at least 50% of all multiple-use or single-use IHAs. Taxa for which >90% of reads originated from a single IHA sample were excluded. By these criteria, a single taxon was enriched in single-use IHAs. Mycoplasma was detected in 3 of the 6 single-use IHAs, and in none of the 8 multiple-use IHAs. Conversely, 7 taxa were enriched in multiple-use IHAs; specifically, Capnocytophaga, Actinomycetales (Order), Granulicatella, Lautropia, Pseudomonas, Eikenella, and Actinomyces. Of note, while Capnocytophaga was detected on 7 of 8 multiple-use and 4 of 6 single-use IHAs, 95.7% of the reads obtained for this taxon originated from multiple-use IHAs, showing a clear enrichment for colonization of multiple-use IHAs. Data for other taxa meeting our enrichment criteria are shown in Fig. 1 and Supplemental data 1. One or more of the red complex genera (Porphyromonas, Treponema, and Tannerella) were detected in 1 out of the 8 multiple-use and in 2 of the 6 single-use IHAs. All of the single-and multiple-use IHAs, excluding one multiple-use IHA, were positive for one or more of the genera in the orange complex bacteria (Fusobacterium, Prevotella and Campylobacter). In total, single-use IHAs corresponded to 35–77.5% of the total reads for each genus identified in the orange complex associated bacteria genera (Supplemental data 1).

Community richness and abundance for each IHA was determined by the Shannon diversity index (H). Within-sample diversity was determined to be comparable between the multiple-use (3.7 ± 0.65) and single-use (3.2 ± 0.75) IHAs (Fig. 2A). Community evenness calculated with Pielou’s evenness index was similar between multiple-use (0.77 ± 0.07) and single-use (0.73 ± 0.08) IHAs (Fig. 2B). These values correspond to comparable community structures in which a single taxon is not dominant. Tighter clustering around the average in the multiple-use IHAs implies similar community representation and corresponds to the mean absolute deviation being lower for multiple-use (0.039) than single-use (0.064) IHAs.

Fig. 2 –

Within-sample diversity (α-diversity) quantified with the Shannon diversity index (A) and Pielou’s evenness (B). Higher average Shannon diversity index was observed in the multiple-use (3.7 ± 0.65), compared to single-use (3.2 ± 0.75) IHAs, although not statistically significant (p > 0.05). Pilou’s evenness score was assessed to determine the community structure with respect to ASV representation. Similar average evenness scores were found for multiple-use (0.77 ± 0.07) and single-use (0.73 ± 0.08) IHAs.

Diversity between samples to quantify the similarity between the individual IHAs was determined using two phylogenetic β-diversity metrics, an unweighted and weighted UniFrac. Clustering by IHA use was not observed in the weighted or unweighted UniFrac, suggesting that usage does not impact overall microbial colonization of the IHAs in terms of composition or abundance (Fig. 3). However, clustering was observed when multiple IHAs were retrieved from a single patient. As described in Table 1, two single-use IHAs (M and N1) were retrieved from patient 12, and two multiple-use IHAs (D1 and E1) were retrieved from patient 2. For the two multiple-use IHAs (D1 and E1) retrieved from patient 2, clustering is observed in both the weighted and unweighted UniFrac (Fig. 3, red circles) indicating that the taxa present are similar, and there are comparable community structures on both IHAs. However, in the single-use IHAs (M and N1) retrieved from patient 12, clustering is observed in the unweighted UniFrac (Fig. 3A, blue stars) but not in the weighted UniFrac (Fig. 3B, blue stars). This indicates that similar taxa are present; however, there are different community structures between these two IHAs. IHAs M and N1 only differ in the presence or absence of Capnocytophaga, Actinomycetales, and Parviomonas, supporting the clustering observed in the unweighted UniFrac. When the relative abundancies are considered, M and N1 show the largest differences in Fusobacterium, Streptococcus, and Neisseria (Fig. 1; Supplemental data 1), which may contribute to the lack of clustering observed in the weighted UniFrac (Fig. 3B).

Fig. 3 –

Diversity between samples (β-diversity) determined with an Unweighted UniFrac (A) and a Weighted UniFrac (B). An Unweighted UniFrac (A) measures diversity by presence or absence of features, and a Weighted UniFrac (B) considers the abundance of the features. Multiple-use IHAs are shown in red and single-use IHAs in blue. Red rings represent two multiple-use IHAs (D1 and E1) retrieved from patient 2. Blue stars represent two single-use IHAs (M and N1) retrieved from patient 12 (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article).

3.2. Surface analysis

OM analysis was performed to assess the degree of morphological changes on IHA surfaces. Observations made on the retrievals were compared to control IHA specimens. As shown in Fig. 4, the control IHAs exhibited a smooth finish with the clear presence of machined lines and strong purple, yellow color on the anodized IHA or silver-colored appearance on the non-anodized IHA. Single and multiple-use IHAs had large amounts of biological debris deposited in the notch area on the IHA head as well as between the screw threads. Single-use IHA B had remnants of blood in the notch and discoloration was observed on the screw area while the majority of the collar region remained unchanged as shown in Fig. 5A, B. Multiple-use IHA E1 showed significant wear, discoloration, and surface abrasion as shown in Fig. 5C, D.

Fig. 4 –

OM images of pristine, unused RC IHA as Control 1: (A) collar neck region (200× magnification) and (B) screw thread (100× magnification). OM images of pristine, unused RN IHA as control 2: (C) notch region (200× magnification) and (D) screw thread (100× magnification).

Fig. 5 –

OM images of RC IHA B after single implantation: (A) collar neck region (100× magnification) and (B) notch region showing remnants of blood (200× magnification). OM images of RC IHA E1 after multiple implantations: (C) collar region showing complete surface discoloration (200× magnification) and (D) collar junction (300× magnification).

SEM analysis was performed to further evaluate morphological changes on the IHA surfaces. All of the control samples evaluated demonstrated smooth machined surfaces with clean screw areas as shown in the representative example of Fig. 7. Single-use and multiple-use IHAs exhibited a greater variety of distinct features on their surfaces that were not present on control IHAs. Single-use IHA J had its collar area peeled off on the edges as shown in Fig. 8A, B. The surface appears rougher and more irregular on the collar and screw areas of the multiple-use IHA E1 Fig. 8C, D. This IHA also had visible pit-like features characteristic of corrosion attack and chipping at the edges of the screw thread region as indicated with arrow marks in Fig. 8C, D.

Fig. 7 –

SEM images of control RC IHA as Control 1: (A) collar junction (91× magnification) and (B) screw thread (91×magnification). SEM images of control RN IHA as control 2: (C) collar junction (49× magnification) and (D) screw thread (126×magnification).

Fig. 8 –

SEM images of IHA J after single implantation: (A) collar region showing signs of surface damage on (127×magnification) and (B) surface delamination (444× magnification). SEM images of IHA E1 after multiple implantations: (C) IHA chipping of the screw thread (584× magnification) and (D) collar showing increased apparent surface roughness (1600×magnification).

The scoring results based on the observations made using optical microscopy (OM) and scanning electron microscopy (SEM) are summarized in Fig. 9. The level of biological contamination and corrosion on the surfaces of IHA tended to increase from control to multiple implantations.

Fig. 9 –

Graphical representation of qualitative scoring of control, single- and multiple-use IHAs.

3.3. Electrochemical analysis

Electrochemical analysis was performed in order to quantitatively evaluate changes in the corrosion behavior of IHA surfaces after single or multiple cycles of exposure to the oral environment. Fig. 10 shows the electrochemical parameters evaluated. No significant difference in Ecorr measurements between control and retrieved IHA specimens were documented. However, the polarization resistance (Rp) was significantly higher (~2 fold) for control IHAs compared to multiple-use IHAs (p < 0.05). The corrosion rate (CR) of both multiple-use and single-use IHAs were significantly higher (~2–3 fold) than the CR for control IHAs (p < 0.05). The Rp and CR results reveal that control IHAs have significantly higher corrosion resistance than single and multiple-use IHAs. Multiple-use IHAs exhibited lower Rp and higher CR on average than single-use IHAs.

Fig. 10 –

(A) Corrosion potential (Ecorr), (B) polarization resistance (Rp), and (C) corrosion rate (CR) of control versus single-use and multiple-use IHAs. *Indicates statistical significance as marked or between all other groups (p < 0.05).

4. Discussion

Implant healing abutments (IHAs) are important temporary components that aid in soft tissue maintenance and can be used to improve the aesthetic outcome of dental implants. Typically, these components are retrieved after early healing, sterilized, and reused on multiple patients [20,36]. The goal of this study was to compare the impact that single and multiple exposures to the oral cavity have on IHAs. We hypothesized that multiple-use IHAs would have lower corrosion resistance and a higher level of surface degradation relative to unused and single-use IHAs and that these surface changes could potentially influence microbial colonization of the IHA surface.

To qualitatively assess the effect of exposure to the oral cavity on the IHA specimens, optical microscopy and scanning electron microscopy were performed. Regardless of usage, visible surface changes were evident on the IHAs following exposure to the oral cavity and were mostly characterized as surface discoloration, pit-like features, and brown spots (Figs. 5, 6, 8, and S1 (Supplemental data 2)). In particular, surface discoloration indicated changes in the native oxidation state of titanium (Ti⁴⁺, TiO₂), with bronze and purple discol-orations characteristic of titanium in its Ti²⁺ (TiO) and Ti³⁺ (Ti₂O₃) states, respectively [37,38]. However, since IHA components are subjected to anodization as a surface treatment, changes in surface coloration could also reflect changes in oxide thickness [39,40]. For instance, a change in abutment color from purple in its pristine form to yellow discoloration suggests a decrease in oxide thickness from >0.04 μm to ~0.03 μm (Figs. 5(C, D) and S2 (Supplemental data 2)) [40]. A thinner oxide layer generally exhibits lower corrosion resistance and hence allows for greater dissolution of metal ions [40]. This surface discoloration was more severe on multiple- use IHAs (Figs. 6 and S2 (Supplemental data 2)) but still present after even a single exposure to the oral cavity (Figs. 5 and S3 (Supplemental data 2)), demonstrating the destructive nature of the oral environment on IHA surfaces. This result was expected, as this component is directly exposed to the fluctuating conditions of the oral cavity, including acidity, which increases the solubility of passive oxide layer and promotes corrosion [41].

Fig. 6 –

OM of RC IHA I after multiple implantations: (A) top surface region showing white deposits in the notch (100×magnification), (B) notch region (300× magnification), (C) collar region showing brown discoloration (300× magnification), and (D) collar junction showing white deposits (100× magnification).

Electrochemical testing was carried out to quantitatively measure the corrosion behavior of the retrieved IHAs. The corrosion potential (Ecorr) is dependent on the thermodynamic stability of the surface oxide, with higher values corresponding to greater stability. The multiple-use IHAs exhibited a decrease in Ecorr value, although not statistically significant, as compared to the single-use and control IHAs (Fig. 10A). The polarization resistance (Rp) quantified the measurement of the resistivity of the oxide layer; a decrease in its value corresponds to a decrease in corrosion resistance. Multiple-use IHAs exhibited significantly lower Rp values when compared to control IHAs (Fig. 10B). Even after one exposure to the oral cavity, the Rp values decreased on average as compared to the control but was not statistically significant. Thus, the trend of decreasing Rp values matched the increasing severity of corrosion damage observed with increasing number of implantations. Conversely, the corrosion rate (CR), which quantified the deterioration rate of the retrieved IHAs as a loss of material mass over time, increased with the number of implantations (Fig. 10C). Moreover, both single- and multiple-use IHAs had significantly higher CR values than control IHA, which further confirmed their greater corrosion susceptibility after exposure to the oral cavity. However, multiple-use IHAs only had a higher average CR value than single-use IHAs. As the Rp and CR values are inversely proportional in theory, observing these expected trends (decreasing Rp values and increasing CR values with the increasing number of exposures) after independently measuring these parameters further validated the corrosion behavior of single- and multiple-use IHAs. Thus, multiple exposures to the oral cavity permanently altered the electrochemical behavior of IHAs by hindering their re-passivation and correlated with the qualitative scoring of corrosion and physical damage on IHAs (Fig. 9).

Surface characteristics and chemistry including porosity, corrosion behavior, roughness, and chemical composition can influence bacterial adhesion on titanium implant surfaces [42]. In turn, bacterial colonization can disrupt the oxide layer and promote titanium corrosion due to metabolic waste products [15,16,41]. Lactic acid is released as a metabolic waste product of Streptococcus, and the decrease in local pH generates the ideal conditions for corrosion [16,43]. Of note, we detected Streptococcus colonization on every IHA analyzed (Fig. 1 and Supplemental data 1). The increase in corrosion rate of the multiple-use IHAs may be a result of the synergistic effect of colonization by bacteria and multiple cycles of sterilization and implantations causing changes to the topography and oxide layer characteristics.

Despite the clear differences in surface characteristics among control, single- and multiple-use IHAs, only few trends related to the number of IHA uses were observed in the taxonomy assessment. While we identified some taxa meeting our criteria for enrichment, clustering by usage was not observed in the β-diversity analysis. Of the taxa identified, Streptococcus was the only genus present on every retrieved IHA and had the second highest relative frequency. The genus Fusobacterium had the highest relative frequency across all IHAs included in microbiome analysis (Fig. 1 and Supplemental data 1). Both of these genera are frequently isolated from healthy and diseased oral cavities [44–46]. Other genera present included Staphylococcus, Granulicatella, Actinomyces, Rothia, Capnocytophaga, Neisseria, and Eikenella, all of these are present in healthy oral cavities [44,47,48]. However, some of these genera (Fusobacterium and Eikenella) include opportunistic pathogens and have been associated with periodontal inflammation and peri-implantitis [46,48]. Additionally, the taxa Prevotella and Filifactor were detected and have been reported to be associated with diseased peri-implant sites [46,49]. However, identification of these genera does not signify disease since pathogenic bacteria are also present in healthy oral cavities [44–46].

The topographical features of an IHA can influence the microbiota present, such that the more similar the surface anatomy, the more similar the microbiome [47]. The lack of significant differences in the α-diversity analysis indicated that similar species richness and abundance is present on single-and multiple-use IHAs, although there was variation within each IHA usage type (Fig. 2). From the β-diversity analysis, a lack of clustering by usage demonstrated that there are additional influencing factors such as individual patient variation. When multiple IHAs were retrieved from an individual patient, the clustering in the unweighted UniFrac indicated similar species were present; this was observed for both single- and multiple-use IHAs (Fig. 3A). Previous studies have demonstrated that a core oral microbiome exists, but there is also a variable microbiome that is individual-specific [47,48,50]. IHA usage may have a role in microbial colonization dynamics; however, individual variation is likely the key factor influencing microbial colonization of IHAs. The lack of clustering in the weighted UniFrac that was observed for the two multiple-use IHAs retrieved from one patient demonstrates a difference in taxa abundance for these IHAs (Fig. 3B). This could be due to different locations of the IHAs in the mouth; however, the lack of additional patient data does not allow us to further explore this hypothesis.

There are limitations to our microbiome analysis. First, while we grouped IHAs by single- and multiple-use, the IHAs evaluated in this study were structurally diverse and consisted of both bone and tissue types, as well as four different connection types (Table 1). Second, the sequencing of only the V1–V3 regions of the 16S rRNA gene did not allow for species-level taxonomy assessment. Third, by not sampling the natural teeth of each patient, differences in the IHA microbiomes due to individual patient variation could not be assessed. Finally, we recovered microbiota using vortexing, but not scraping or sonication, meaning that we may not have recovered the entirety of the biofilm colonizing the IHA surface.

What is the potential impact of corrosion and surface damage on IHA functionality and clinical use? When an IHA is in function, the different regions of the device are exposed to various atmospheric conditions, such as the head which is directly exposed to an aerobic environment while the screw can experience a microaerophilic and potentially an anaerobic environment. Microgaps between IHAs and the hollow component of the subgingival screw act as potential reservoirs for pathogenic bacterial species [52]. Increased corrosion and wear may influence microgap sizes, as well as bacterial adhesion. Corrosion abrasions and chipping were observed on the threads of the IHAs, and biologic debris was identified on the screw portion of the IHAs (Figs. 5, 6, and 8). These findings indicate biologic leakage and colonization of the portion of the IHA that would be under microaerophilic or anaerobic atmospheric conditions. Overall, the practice of multiple implantations may change the surface morphological and composition features of IHAs due to the accumulation of biological deposits, corrosion products and may increase the bacterial load on their surfaces. Thus, clinical reuse of IHAs may play a vital role in the contribution of bacterial colonization at the implant-IHA interface, which may adversely affect the long-term performance of implant systems. Ultimately, longitudinal studies of patients that receive both single and multiple use IHAs should be performed to determine whether negative outcomes (peri-implant mucositis; peri-implantitis) significantly differ between the two IHA modalities.

Keeping limitations in mind, future studies should characterize the change in microgap size between single- and multiple-use IHAs, as well as identify whether different micro-biota are present at the regions of the IHA under different atmospheric pressures. Tests using IHAs for which the exact number of uses is known should be carried out in order to determine if frequency and duration of oral cavity exposure and corrosion are directly correlated. Additionally, studies examining only one IHA type and junction would improve the understanding of the effects of multiple exposures to the oral cavity. A study examining two IHAs a single- and multiple-use retrieved from one patient would provide valuable information and aid in decreasing the influence of individual patient variation.

Uncited references

[51].