Circulating monocytes from aseptic loosening patients exhibit enhanced tropism toward periprosthetic tissues in NOD-SCID mouse model

Abstract

Aseptic loosening (AL) remains the leading cause of implant failure after arthroplasty and is driven by chronic periprosthetic inflammation induced by particulate wear debris. Circulating monocytes contribute to this process by homing to inflamed periprosthetic tissues and propagating osteolytic inflammation. However, it remains unclear whether their enhanced recruitment reflects local chemoattractant signals alone or a state of systemic immune priming. To investigate the homing and migratory capacity of circulating monocytes from patients with aseptic loosening toward inflammatory periprosthetic tissues using a NOD-SCID xenograft model. Revision periprosthetic tissues (RPT) from AL patients and normal synovial tissues (NST) from anterior cruciate ligament reconstruction patients were implanted subcutaneously into contralateral quadriceps regions of NOD-SCID mice. Unrelated human peripheral blood monocytes (n = 6 each) were isolated from healthy volunteers (G1), primary arthroplasty patients (G2), and aseptic-loosened revision patients (G3), fluorescently labelled, and intravenously injected into mice. In vivo monocyte trafficking was quantified by serial fluorescence imaging at 1, 24, and 72 h. Monocyte infiltration into RPT was significantly greater than into control tissues across all groups. Recruitment was highest in G3, followed by G2 and G1 at 72 h (p < 0.05), with early migration evident by 24 h in all groups except G1. Multivariate regression identified patient group and implant duration as independent predictors of monocyte migration (R²=0.641, p < 0.001). Periprosthetic tissues are intrinsically chemotactic to circulating monocytes; however, monocytes from AL patients exhibit a markedly enhanced, hyper-responsive migratory phenotype consistent with systemic immune priming. These findings identify monocyte reprogramming as a previously underappreciated contributor to implant failure and suggest that targeting systemic innate immune activation may enable earlier risk stratification and novel therapeutic strategies to prevent or delay aseptic loosening.

Introduction

Joint arthroplasty registries have reported that implant failures are predominantly caused by aseptic loosening (AL). This accounts for 30–60% of all knee and hip revision surgeries^1–3. AL is primarily driven by the production of wear debris particles at the periprosthetic tissue interface, which triggers local inflammatory cascades^4–6. Wear particles are phagocytosed by macrophages, which later release pro-inflammatory mediators as well as damage-associated molecular patterns (DAMPs) that further activate other macrophages and neighbouring cells. These include neutrophils, fibroblasts, and endothelial cells, contributing to a destructive pro-inflammatory environment^7,8. These inflammatory factors, therefore, promote the recruitment of immune cells and other precursor cells, including osteoclast precursors⁹. As the disease progresses, the formation of osteoclasts carrying out bone resorption activities at the bone-implant interface, referred to as periprosthetic osteolysis, becomes greater. This would eventually lead to implant loosening and failure^10,11.

Periprosthetic tissue environments, which are loaded with chemoattractant cues, play a crucial role in various functions, including phagocytosis, the release of pro-inflammatory cytokines and chemokines, immune cell trafficking, and tissue infiltration^12,13. Previous studies have explored the cytokine profiling of periprosthetic tissue derived from aseptic loosened-revision arthroplasty and reported a significant elevation of pro-inflammatory cytokines^14–16. The release of chemokines, which generally functions to facilitate the migration and activation of immune cells such as monocytes, is reported to be associated with the progression of aseptic loosening¹⁷.

Monocytes, which generally reside in the circulatory system, carry out their immune surveillance role and are recruited into specific tissue regions following chemotaxis gradients in cases of inflammation¹⁸. Chemokines facilitate the monocyte trafficking process and further infiltration into the inflamed tissue niche^19,20. Monocytes then get activated and further differentiate into pro-inflammatory macrophages^21,22. Studies have reported that migrated monocytes can differentiate into osteoclasts, which are key players in the periprosthetic osteolysis process^23,24.

Recent studies also suggest that patients exposed to chronic inflammation due to aseptic loosening may undergo a process called “priming” that promotes phenotypic changes, thereby intensifying their responsiveness towards subsequent chemoattractant stimuli^25,26. The priming process is a rapid, short-term pre-activation of immune cells that occurs in the peripheral blood or bone marrow, maintaining cells in a “state of readiness” to respond to environmental challenges, including subsequent pro-inflammatory exposure²⁷. Prolonged chronic inflammation exposure during AL progression may influence monocyte behaviour and its migratory capacities; however, this remains poorly understood. We hypothesized that the prolonged inflammatory exposure promotes “monocytes priming”, which allows for rapid, exaggerated monocytes migratory responses to the periprosthetic tissue stimuli.

The comprehensive understanding of the migratory capacity of primed monocytes following prolonged inflammatory exposure in aseptic loosening progression requires further investigation. The findings may provide valuable insights into the dynamics of disease progression mechanisms and pave the way to a potential therapeutic target. Nevertheless, to mimic the complexity of the human immune system in laboratory settings remains challenging. Therefore, an immunodeficient (NOD-SCID) murine model was particularly utilized in this in vivo study, which offers a useful platform to study the human monocyte migration in a controlled environment without risking xenogeneic rejection would be ideal. Techniques involving human periprosthetic tissue transplantation in these murine models allow the restoration of the inflamed tissue microenvironment that enables real-time tracking of fluorescently labelled human monocytes.

This present study was thus conducted using such model to investigate the effect of prolonged, chronic inflammatory exposure on the migratory capacity of monocytes derived from aseptic loosened revision arthroplasty patients. Human revision aseptic loosened periprosthetic tissue transplantation was transplanted in this model, followed by an intravenous injection of fluorescently labelled monocytes isolated from revision arthroplasty patients. The tracking of injected monocytes enables us to examine their migration dynamics toward transplanted revision periprosthetic tissue under controlled conditions. This approach aims to elucidate the affinity of monocyte recruitment in peri-prosthetic tissue during disease progression, thereby indirectly demonstrating that a “priming” effect may be the underlying reason for higher monocyte migration and therefore accelerated implant loosening.

Methods

Human sample collection

The human tissue samples and retrospective data collection in this study were approved by the University of Malaya Medical Centre-Medical Research Ethics Committee (UMMC-MREC) (Ethics No: 20211029-10716), and informed consent was obtained from all patients. All procedures involving human tissue samples were carried out in accordance with ethical standards and institutional guidelines, in line with the principles of the Declaration of Helsinki. Revision periprosthetic tissues (RPT) were collected from patients (n = 6) undergoing revision arthroplasty, whilst normal synovial tissues (NST) were obtained from patients (n = 6) undergoing anterior cruciate ligament (ACL) reconstruction surgery at University Malaya Medical Centre (UMMC). Peripheral blood samples not related to tissue donors were also collected from stable primary arthroplasty patients, i.e., patients going for primary knee arthroplasty (n = 6; mean age = 81) and aseptic-loosened revision arthroplasty patients (n = 6; mean age = 78) attending annual follow-up at the orthopaedics clinic, and healthy volunteers (n = 6; mean age = 29) were recruited as a control; physiological baseline to establish normative range of monocyte chemotaxis.

Ethics approval and animal care

The use of the murine model in this in vivo study was approved by the Institutional Animal Care and Use Committee (IACUC), University of Malaya (Ethics No: 2020–231203/ORTHO/R/KG). All animal experimental procedures involved complied with the institutional guidelines and ARRIVE guidelines. Before any procedure, anaesthesia agents were used, and post-operative care was provided to minimise pain and infection risks. At the end of the experiments, mice were humanely euthanized by cervical dislocation under deep anaesthesia. All measures were taken to minimize the animal suffering and discomfort throughout in vivo experiments.

Tissue transplantation in NOD-SCID mice

This study utilized the use of twenty-four male (n = 24), aged between 12 and 14 weeks, severe combined immunodeficient (NOD-SCID; Nomura Siam International Co., Ltd., Thailand) mice model. Before the transplantation procedure, mice were anesthetized with Xylazine (10 mg/kg) and Ketamine (50 mg/kg) and then placed in a prone position. The lumbar-sacral region was shaved, treated with povidone-iodine, and draped with a surgical towel, exposing the area of interest. Under sterile conditions, a 3 mm x 3 mm piece of human RPT was transplanted on the small incision made at the right lumbar-sacral region, while human NST was transplanted at the left area. Incision sites were secured using 6 − 0 silk suture. For postoperative care, the transplanted mice were exposed to a heat lamp for 15 min and received painkillers (Meloxicam, 5 mg/kg) and antibiotics (Kombitrim, 10 mg/kg) via intramuscular injection.

Isolation and PKH26-fluorescence labelling of peripheral blood monocytes

Peripheral blood samples (n = 18) were collected in citrate blood tubes and isolated using the density centrifugation method. Briefly, samples were diluted with phosphate-buffered saline PBS) and layered over Histopaque separation media (1:1 ratio). Following centrifugation (2000 rpm, zero brake, 20 min), the mononuclear cell layer containing monocytes was collected. The isolated peripheral blood (PB)-derived monocytes were used for cellular labelling with a fluorescent dye (PKH26 Red Fluorescent Cell Linker Kit, Sigma-Aldrich, USA). This assay was performed by following the manufacturer’s protocol (Sigma-Aldrich Co. LLC, USA). PKH26-monocytes obtained were resuspended in PBS and stored in a chiller box for further use.

Intravenous injection of PKH26-monocytes into transplanted NOD-SCID mice

After 72 h post-transplantation, PKH26-monocyte injection was performed, with saline injection serving as the control. Before the injection procedure, mice were anesthetized with Xylazine (10 mg/kg) and Ketamine (50 mg/kg). PKH26-monocytes that were freshly prepared and stored in a chiller box were then thawed. Intravenous injection of PKH26-monocytes (2 × 10^6) through the tail vein was performed using a needle size of 27G.

In vivo live imaging

The injected PKH26-monocytes were tracked non-invasively using In Vivo Multispectral Fluorescence FX PRO (MS FX PRO Bruker System, USA). Mice were placed into a small animal chamber located inside the machine. The Region of Interests (ROIs) were specifically created on the transplanted (NST/RPT) sites to ensure systemic, repeatable imaging across all groups. Fluorescence intensity was normalized against baseline measurements using saline-injected mice (control) to account for tissue autofluorescence, ensuring that all reported metrics represent stimulus-driven recruitment. By utilizing these rigorous ROIs and normalization criteria, the quantification of monocyte accumulation was standardized, thereby minimizing experimental variability and ensuring reliability of the observed migratory kinetics. Imaging procedures were carried out at 1 h, 24 h, and 72 h. Image acquisition and data visualization of the emitted fluorescent signals were performed using Carestream Molecular Imaging software, with regions of interest (ROIs) explicitly defined on the transplanted tissue regions.

Statistical analysis

In vivo experiments were conducted using six independent samples collected from different donors within each experimental group. Data were represented as mean ± standard deviation (SD). Statistical significance was performed using Student’s t-test and one-way analysis of variance (ANOVA), followed by Bonferroni post-hoc multiple comparison. A 95% confidence level (p < 0.05) was selected to determine statistical significance in the datasets. Statistical analysis was performed using IBM SPSS Statistics version 29 software (SPSS Inc., Chicago, Illinois, USA). Pearson correlation, nonlinear, and multiple regression analyses were conducted using GraphPad Prism version 10 software (GraphPad Software, LLC., USA).

Results

Demographic profile of patient cohorts

Human peripheral blood-derived monocytes were collected from a total of 18 patients, divided equally into three groups: healthy volunteers (G1), stable primary arthroplasty patients (G2), and revision arthroplasty patients (G3). G1 consisted of healthy individuals with a mean age of 28.67 ± 3.27 years, who had not undergone arthroplasty surgery and served as the negative control group. G2 consisted of patients with a mean age of 81.17 ± 6.77 years who had undergone primary arthroplasty and were considered to have stable implants, with a reported duration of 5.17 ± 0.65 years. G3 consisted of patients with a mean age of 77.83 ± 3.58 years who had undergone primary arthroplasty, with a history of first-time implantation for 15.67 ± 1.89 years, and were diagnosed with implant failure due to aseptic loosening. These patients were being planned for revision arthroplasty. Detailed demographic characteristics of these patients are presented in Table 1.

Table 1.

Descriptive data of demographic characteristics for every group cohort.

	Healthy donor (N = 6)	Stable primary arthroplasty (N = 6)	Revision arthroplasty (N = 6)	Adjusted p-value
Demographic characteristics
Age in years	28.67 ± 3.27	81.17 ± 6.77	77.83 ± 3.58	< 0.0001	****
Arthroplasty surgery	No	Yes	Yes
Implant duration in years	-	5.17 ± 0.65	15.67 ± 1.89	0.0004	***

PKH26-monocyte injection in the transplanted NOD-SCID mice model

To investigate the migration capacity of monocytes in an in vivo model, PKH26-monocytes isolated from different patient cohorts were injected into a transplanted NOD-SCID mouse model, and their movement was tracked based on fluorescence emission detection, with saline injection serving as a control. Four groups were compared, as outlined in Table 2.

Table 2.

Different groups for post-transplantation cell injection.

Group	Labelled	PB-Monocyte source for cell injection
1	G0	X (saline only)
2	G1	Healthy volunteers
3	G2	Stable primary arthroplasty
4	G3	Revision arthroplasty

Descriptive analysis of mean fluorescence intensity across groups, transplanted tissue types, and timepoints

Fluorescence signals detected in the ROI regions were recorded, and descriptive analysis was performed using the mean intensity across all groups, transplanted tissue types, and time points. The descriptive data consisted of mean, standard deviation (SD), standard error (SE), median, and coefficient of variation (CV). It was observed that there was a variation in the mean intensity values, with detailed statistics presented in Table 3.

Table 3.

Descriptive data of mean fluorescent intensity detected in explant regions across all groups and timepoints. T0 corresponds to 1 h from monocyte injections, whereas T1 and T2 correspond to 24 and 72 h, respectively.

Group	Explant	Timepoint	Mean	St. Deviation	St. Error	Median	CV (%)
G0	NST	T0	8.355	0.602	0.246	8.352	6.573
		T1	8.355	0.602	0.246	8.352	6.573
		T2	8.317	0.737	0.301	8.147	8.088
	RPT	T0	8.386	0.255	0.104	8.394	2.772
		T1	8.386	0.255	0.104	8.394	2.772
		T2	8.779	0.865	0.353	8.779	8.995
G1	NST	T0	9.692	1.000	0.408	9.947	9.415
		T1	11.981	1.317	0.538	11.711	10.038
		T2	13.183	1.518	0.620	13.721	10.514
	RPT	T0	9.954	1.213	0.495	9.879	11.124
		T1	13.696	1.463	0.597	13.696	9.750
		T2	16.196	0.846	0.345	16.312	4.769
G2	NST	T0	9.979	0.792	0.323	10.074	7.249
		T1	12.244	1.703	0.695	12.750	12.695
		T2	13.850	1.613	0.659	13.796	10.632
	RPT	T0	10.187	1.051	0.429	10.036	9.419
		T1	15.333	1.515	0.619	15.502	9.022
		T2	19.735	1.682	0.687	19.790	7.779
G3	NST	T0	9.158	1.303	0.532	9.307	12.992
		T1	10.069	1.253	0.511	10.214	11.358
		T2	11.612	0.748	0.305	11.563	5.879
	RPT	T0	9.107	1.794	0.732	9.306	17.983
		T1	14.083	2.160	0.882	13.585	14.002
		T2	22.105	1.783	0.728	22.739	7.363

Significant PKH26-monocyte migration into the transplanted RPT region

Fluorescence signal intensity, as an indication of PKH26-monocyte migration intensity, was compared across transplanted NST and RPT regions (Fig. 1). At 1-hour post-injection (T0), no significant difference was observed across all groups (G0-G3). At 24 h after injection (T1), G2 exhibited a significantly higher migration of PKH26-monocytes into RPT (15.333 ± 1.515) as compared to NST (12.244 ± 1.703) with a p-value of 0.0078. In G3, PKH26-monocyte migration into RPT (14.083 ± 2.160) was also significantly higher than NST (10.069 ± 1.253; p = 0.0028). For 72 h post-injection (T2), significant differences in migrated PKH26-monocytes into NST and RPT were observed across all groups, except for G0. For G1, higher PKH26-monocyte migration in RPT (16.196 ± 0.846) than NST (13.183 ± 1.518; p = 0.0017). Migrated PKH26-monocytes in G2 showed higher RPT (19.735 ± 1.682) compared to NST (13.850 ± 1.613; p = 0.0001). G3 exhibited the highest migration of PKH26-monocytes into RPT (22.105 ± 1.783), significantly compared to NST (11.612 ± 0.748; p < 0.0001).

Fig. 1.

Fluorescence intensity comparison between NST and RPT, across all groups and timepoints. Data represented as mean values indicative of PKH26-monocyte migration intensity. Significant differences between tissues within each group and time point are indicated by asterisks (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001). Error bar = ± SD.

Revision arthroplasty cohort exhibited greater PKH26-monocytes migration

The migration intensity into transplanted tissues was compared across all groups (Fig. 2). It was observed that there was no significant difference at 1 h (T0) and 24 h (T1) post-injection. PKH26-monocyte migration into the RPT region was highest in G3 (22.11 ± 1.78), followed by G2 (19.735 ± 1.68) and G1 (16.196 ± 0.85) at 72 h (T2). There was no significant difference in PKH26-monocyte migration into NST across all groups and timepoints. Fold changes were also calculated using G0 as the baseline reference. It was revealed that G1 exhibited a 0.85-fold increase, followed by G2 with a 1.26-fold increase, and G3 with a 1.54-fold increase, which was significantly higher than that of G2 and G3 (p < 0.0001). Non-linear regression analysis was conducted for T2; an exponential growth pattern was observed in PKH26-monocyte migration into the RPT region across the group, as indicated by a significant p-value (p < 0.0001) and a strong model fit with an R² value of 0.8432.

Fig. 2.

(A) Fluorescence intensity and fold changes across all groups, transplanted tissues, and timepoints—data represented as mean values indicative of PKH26-monocyte migration intensity. Significant difference between groups is indicated by asterisks (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001). Error bar = ± SD. (B) Non-linear regression analysis of fold change in T2. The analysis reveals a significant exponential increase in the PKH26-monocyte migration rate into RPT, from G1 to G3 (p < 0.0001, r² = 0.8432).

PKH26-monocytes migration into the RPT region increases over time

Migration rates of PKH26-monocytes into transplanted NST and RPT regions were observed at three different time points: 1 h (T0), 24 h (T1), and 72 h (T2) post-injection. There was no significant difference in PKH26-monocyte migration into NST across all groups and time points. Fold changes were calculated across time points in every transplanted tissue and group, using T0 as the baseline reference. There was a significant increase in PKH26-monocytes migrating into RPT 72 h after injection, shown in all groups except G0. In G1, PKH26-monocytes migrated into RPT, exhibiting a 0.64-fold increase at T2; a significant increase was observed between 24 h (T1) and 72 h (T2) post-injection (p = 0.0046). For G2, a similar trend was observed: PKH26-monocyte migration into RPT continuously increased until T2 (0.96-fold), which was significantly higher than T1 (p < 0.0001). G3 demonstrated the highest increase in PKH26-monocyte migration into RPT, with a 1.48-fold increase at T2, substantially higher than at T0 and T1, respectively (p < 0.0001). In contrast, no significant difference was observed in the number of PKH26-monocytes migrating into NST across all timepoints and groups (see Fig. 3).

Fig. 3.

Fluorescence intensity and fold changes comparison across timepoints within each group and transplanted tissue—data represented as mean values indicative of PKH26-monocyte migration intensity. Significant differences between tissues within each group and time point are indicated by asterisks (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001). Error bar = ± SD.

Nonlinear regression analysis of PKH26 migration rate into RPT tissue

To examine the relationship between patients’ groups/cohorts, timepoints, and PKH26 migration rates into transplanted RPT tissue specifically, non-linear regression was performed (Fig. 4). It was observed that the goodness of fit, as measured by the coefficient of determination (R²), varied across groups. The R² values were 0.09, 0.45, and 0.71 for G1, G2, and G3, respectively. G3 exhibited a strong fit, with 71% of the variation in the PKH26 Migration rate, demonstrating a robust relationship modelled by the nonlinear regression. These results, therefore, highlight how PKH26-monocyte migration into RPT tissue was regulated across groups and time points.

Fig. 4.

(A) Fluorescence intensity and fold changes comparison of migrated PKH26-monocytes into the RPT region across timepoints and groups—data represented as mean values indicative of PKH26-monocyte migration intensity. Significant differences between each group and time point are indicated by asterisks (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001). Error bar = ± SD. (B) Non-linear regression analysis of different groups and different timepoints influencing fluorescence intensity outcome.

Multiple linear regression model for PKH26-monocytes migration rate

To assess the unique contribution of each variable to the PKH26-monocytes migration, Pearson correlation and exploratory multiple linear regression analysis were performed (Fig. 5). Variables including patients’ groups/cohorts, age, transplanted tissue types, and implant status, as well as duration post-arthroplasty, with fluorescence intensity as the dependent variable indicative of PKH26-monocyte migration. Pearson correlation analysis demonstrated a significant positive correlation between group (r = 0.23), tissue types (r = 0.75), age (r = 0.26), arthroplasty status (r = 0.26), and implant duration (r = 0.14) with the dependent variable, PKH26-monocyte migration rate.

Fig. 5.

(A) Pearson correlation matrix. (B) Normal Probability-Probability (P-P) plot of the standardized residuals regression model for the dependent variable (fluorescence intensity). The points shown were closely following a diagonal line, indicative of normal distribution, thus supporting the suitability of the regression model.

The outcome of the multiple linear regression was statistically significant, indicating that these factors collectively reflected a meaningful relationship with the PKH26-monocyte migration rate (adjusted r² = 0.582, p < 0.0001).

The equation for the regression model was as follows:

Based on the standardized residuals (P-P plot), it showed that these residuals were approximately normally distributed, supporting the validation of the regression model assumptions (Fig. 4). Multicollinearity diagnostics revealed elevated VIF values for several covariates; the model should be interpreted cautiously as exploratory. These results indicate that patient cohorts, implant status/duration, and age were individually associated with PKH26-monocyte migration, with tissue type showing the strongest association.

Discussion

The present study provides novel evidence that circulatory monocytes derived from aseptically loosened revision arthroplasty patients exhibited heightened migratory capacity, suggesting that these cells were primed for activation and rapidly responded to subsequent chemoattractant cues following prolonged inflammatory exposure. As one of the key players during inflammation, monocytes become activated and rapidly migrate into the periprosthetic tissue niche during the aseptic loosening progression^28–30. Over time, the progressive elevation in monocyte migration intensity demonstrated across patient cohorts—from healthy volunteers through stable post-arthroplasty to revision arthroplasty patients—exhibits a clear association between duration and intensity of inflammatory exposure and monocyte responsiveness towards chemoattractant.

Monocyte migration plays a crucial role, representing a critical determinant of inflammation intensity and tissue damage in the periprosthetic tissue microenvironment. The efficiency of the monocyte migration and further tissue infiltration directly influences the extent of local inflammation and osteolysis progression rate⁸. The recruitment process of circulatory monocytes into the inflammation site is highly facilitated by chemoattractant gradients, adhesion molecule expression, and tissue-derived inflammatory stimuli²¹. Our findings demonstrated that the migratory capacity of circulatory monocytes into transplanted periprosthetic tissue was significantly higher compared to normal synovial tissue across G1, G2, and G3 over time. At 72 h, monocytes from revision arthroplasty patients exhibited nearly twice the migration value into the RPT region (22.105 ± 1.783) compared to the NST region (11.612 ± 0.748). This preferential migration may suggest that the periprosthetic tissue may contain a chemoattractant that draws monocytes to migrate to this specific region.

The inflammatory cytokines and chemokines play a crucial role in facilitating the activation, recruitment, and subsequent infiltration of immune cells into tissues during inflammatory events^31,32. Although this study did not thoroughly screen for the inflammatory cytokines’ presence, few studies have examined the periprosthetic tissues collected from aseptic-loosened revision arthroplasty patients^33–35. They reported elevated levels of various cytokines, including interleukin-1-beta (IL-1B), interleukin-6 (IL-6), interleukin-8 (IL-8), interleukin-10 (IL-10), tumor necrosis factor alpha (TNF-α), and several chemokines such as monocyte chemoattractant protein-1 (MCP-1) and macrophage inflammatory protein-1 (MIP-1) were observed^36,37. These cytokines were predominantly expressed by the resident fibroblast, vascular endothelial cells, and mainly macrophages, which contribute to the chronic inflammatory cascades at the bone-implant interface^33,34,37.

The notable migratory capacity into the periprosthetic tissue regions exhibited by circulatory monocytes derived from aseptically loosened revision arthroplasty patients (1.54-fold) into the transplanted RPT region was significantly higher than both stable post-arthroplasty patients (1.26-fold) and healthy individuals (0.85-fold), as compared to saline controls at 72 h post-injection. This distinguished migratory capacity may suggest that monocytes were primed for activation and rapid responsiveness towards subsequent inflammatory stimuli, owing to their prior activation or priming effect in response to prolonged inflammatory exposure during the progression of aseptic loosening. Studies have reported that monocyte activation initially occurs in the bone marrow before recruitment, a process referred to as priming. Recent studies have demonstrated that monocytes can develop a long-lasting functional reprogramming following a persistent inflammatory exposure, which heightened their responsiveness and migratory capacities during disease progression³⁸. Monocyte priming results in phenotypic alterations due to chronic low-grade inflammatory exposure, which was also reported in other pathogeneses such as atherosclerosis, rheumatic arthritis, and chronic liver diseases³⁹.

Generally, monocytes are precursors that develop in the adult bone marrow and are recruited through the circulation to replenish tissue-resident macrophages in the event of tissue injury or inflammation. Monocytes are highly dependent on chemokines such as C-C motif chemokine ligand 2 (CCL2), C-C motif chemokine ligand 5 (CCL5), and C-C motif chemokine ligand 2 (CCL7) for this process^38,39. It was reported that bone marrow stromal cells produce CCL2 in response to circulating toll-like receptor ligands, thereby contributing to the activation of precursor cells, including monocytes³⁹. A previous study reported that monocyte culture exposed to wear particles showed elevated CCL2/MCP-1 levels, and significant migration was also reported post-exposure³⁵. Other findings reported that CCL2 is required to activate C-C motif Chemokine receptor 2 (CCR2) in monocytes, therefore contributing to their egress from bone marrow⁴⁰. During inflammation, priming is necessary not only to activate monocytes for recruitment, but it is also required to enhance their responsiveness towards inflammatory stimuli once they reach the tissue site^41,42. The continuous priming and recruitment of monocytes may also exacerbate chronic inflammation during the progression of aseptic loosening. Given the significant presence of CCL-2/MCP-1 in periprosthetic tissue and its crucial role in facilitating the trafficking of immune cells, specifically monocytes, the CCL2-mediated signalling mechanism may be highly involved in promoting the migratory capacity of circulating monocytes^15,35. While a mechanistic investigation into the CCL2/CCR2 signalling pathway was not performed, our findings indicate that this heightened responsiveness is a functional outcome of the loosening-prone state. Whether these migratory behaviours are explicitly driven by the CCL2/CCR2 axis remains hypothetical; therefore warrants future investigation into the specific signalling cascades involved.

The correlation and multiple regression analysis also revealed that host factors, including patients’ cohort, tissue type, age, arthroplasty status, and implant duration, collectively associated with the PKH26-monocytes migration capacity (r² = 0.641, p < 0.001). These findings confirm that the inflammatory characteristics of periprosthetic tissue and the prolonged aseptic loosening exposure in revision arthroplasty patients are the primary driver that orchestrates monocyte migratory capacity and tissue infiltration functions. Patients with longer implant durations post arthroplasty who remain stable may represent a cohort with inherently lower inflammatory response, whereas those aseptic loosened revision patients exhibited more aggressive inflammatory, primed phenotype due to the heightened responsiveness. However, a comprehensive, detailed mechanism that dictates monocyte priming and phenotypic changes during aseptic loosening progression following arthroplasty warrants further investigations.

In this study, fluorescent labelling of the cellular membrane using PKH26 dye was employed to assess monocyte trafficking into transplanted periprosthetic tissue in the NOD-SCID mice model. PKH26 labelling is an established technique used to track cells in vitro and in vivo⁴³. It is a long-chain carbocyanine, a lipophilic dye that binds to the exposed lipid bilayer of cells through its aliphatic tails, forming a strong noncovalent bond and thereby providing long-term retention and stable fluorescence signals⁴⁴. PKH26 labelling involves an easy and rapid staining procedure, which can be detected under fluorescent microscopy or an in vivo live imaging system. Our study clearly demonstrated a significant increase in fluorescent signal retention in transplanted NOD-SCID mice, specifically in the periprosthetic tissue region following cell injection. The use of an immunocompromised, nude mice model (NOD-SCID) as a host plays a crucial role in investigating the dynamics of immune cell interaction with inflammatory stimuli exposure released from the tissue niche⁴⁵. Their impaired innate and adaptive immunity, due to the dysfunction of functional T and B lymphocytes, makes them an ideal tool for studying the complexity of the human immune system⁴⁶. Overall, the experimental techniques employed in this study enabled a quantitative assessment of the migratory capacity of circulatory monocytes under physiologically relevant conditions, thereby providing robust mechanistic insights into the possible effect of priming in aseptic loosening pathogenesis.

This study highlights that monocytes derived from revision arthroplasty patients with long-term arthroplasty implants possessed heightened migratory capacity, therefore providing a novel framework for clinical risk stratification. From a clinical perspective, the alterations in monocyte phenotype and functional outcomes could potentially be used as an early indicator to help identify high-risk patients who have the onset of loosening progression, which often goes undetected under radiographic evidence. Monocyte priming process and further functional outcomes can serve as the point of intervention to help delay the inflammation cascades leading to progressive osteolysis. While systemic immunosuppression is not feasible for arthroplasty patients, targeted inhibitors – such as a CCR2 antagonist – could theoretically blunt the ‘primed’ monocytes phenotype and decelerate recruitment that may contribute to bone loss in the periprosthetic tissue region in identified high-risk patients. It is important to emphasize that these implications remain exploratory; future prospective studies are necessary to establish monocytes migratory velocity correlations with radiographic metrics of implant failure and to define the longitudinal trajectory of monocytes function post-arthroplasty.

However, despite the robust nature of the study, several limitations need to be highlighted and should be considered in the interpretation of our findings. Small sample size (n = 6/group) due to recruitment constraint limits the statistical power; therefore, replication in larger cohorts may be warranted. Given the significant role of periprosthetic tissue in determining the migratory rate of circulatory monocytes, a detailed tissue profiling and cellular characterization would be necessary. The absence of extensive tissue analysis post-migration also limits our understanding of cellular dynamics. While this investigation provides compelling evidence of monocyte priming in aseptic loosening, the use of immunocompromised nude mice limits full recapitulation of human immune interactions. Species differences in chemokine signalling and tissue remodelling may influence monocyte trafficking patterns observed in the xenograft model. The study demonstrates enhanced migration but does not elucidate underlying molecular pathways. Specific priming mechanisms—such as upregulated chemokine receptors (CCR2, CXCR4), other possible related adhesion molecules, or specific transcription factors—remain uncharacterized, limiting insights into therapeutic targets. Therefore, whilst the study does suggest priming as the cause for enhanced monocyte migration in AL patients, there is a need for more robust models to be conducted for conclusive validation.

It would therefore be prudent for future studies to utilize humanized mouse models with reconstituted immune compartments to more accurately mimic clinical aseptic loosening. Single-cell RNA sequencing of primed monocytes could map transcriptional changes driving enhanced migration, identifying novel biomarkers and therapeutic windows. Longitudinal tracking of monocyte differentiation into osteoclasts within revision tissue grafts would link acute infiltration to chronic osteolysis. In addition, incorporating advanced imaging (e.g., intravital two-photon microscopy) that enables real-time visualization of monocyte-endothelial interactions and tissue penetration dynamics would be preferred. Another aspect would be in expanding patient cohorts with comprehensive phenotyping (cytokine profiles, genetic polymorphisms in chemokine pathways) that would further refine predictive models for revision risk. This would then allow clinical trials targeting primed monocyte pathways—such as CCR2 antagonists or NF-κB inhibitors— to be conducted and thus validate therapeutic efficacy in preventing aseptic loosening progression in patients.

Conclusion

In summary, RPT tissue significantly promotes monocyte migration and tissue infiltration across all patient cohorts. Monocytes from patients with aseptic loosening exhibit a heightened migratory affinity for RPT, an observation consistent with functional priming driven by the local disease progression. Our findings suggest that prolonged inflammation in aseptic loosening creates a priming environment that favours monocyte activation, which enhances monocyte responsiveness to chemoattractant stimuli present in periprosthetic tissues. However, the absence of any significant migration into NST across all groups over time indicates that priming alone is insufficient to promote monocyte migration. Furthermore, patients with longer durations of implant arthroplasty, particularly those with advancing age, are more likely to develop monocytes that are attracted to RPT, thereby increasing the risk of aseptic loosening. While these observations suggest that monocytes priming and a heightened migratory phenotype are key features of loosening-prone states, future molecular studies are needed to confirm the precise signalling mechanisms involved. Collectively, these exploratory insights provide a foundation for developing risk-stratification strategies and early intervention points for aseptic loosening.

Abbreviations

AL

Aseptic Loosening

NOD-SCID

Non-obese Diabetic/Severe Combined Immunodeficiency

NST

Normal Synovial Tissue

DAMPs

Damage-Associated Molecular Patterns

UMMC-MREC

University of Malaya Medical Centre-Medical Research Ethics Committee

ACL

Anterior Cruciate Ligament

IACUC

Institutional Animal Care and Use Committee

PB

Peripheral Blood

SFM

Serum Free Media

RPMI-1640

Roswell Park Memorial Institute 1640 medium

PBS

Phosphate buffered solution

SD

Standard Deviation

ANOVA

Analysis of Variance

SE

Standard Error

CV

Coefficient of Variation

IL-1B

Interleukin-1-Beta

IL-6

Interleukin-6

IL-8

Interleukin-8

IL-10

Interleukin-10

TNF-α

Tumor Necrosis Factor-alpha

MCP-1

Monocyte Chemoattractant Protein-1

MIP-1

Macrophage Inflammatory Protein-1

CCL2

C-C motif Chemokine Ligand 2

CCL5

C-C motif Chemokine Ligand 5

CCL7

C-C motif Chemokine Ligand 7

CCR2

C-C motif Chemokine Receptor 2

CCR4

C-C Motif Chemokine Receptor 4

RNA

Ribonucleic Acid

NF-κB

Nuclear factor kappa-B

Author contributions

All authors meet ICMJE criteria. [N.A.] performed the in vitro and in vivo experiments, data acquisition, statistical analyses, and drafted the initial manuscript. [H.F.] was involved in the in vitro optimization and data acquisition. [A.Z.] was involved in animal handling, tissue transplantation, preparation, and cell injection for in vivo. [K.G.] designed the experimental protocols and supervised all experimental procedures. [K.A.A] provided funding acquisition, patient recruitment, and clinical expertise. [T.K.] supervised the entire project, reviewed data analysis, and approved the final manuscript version.

Funding

This study was funded by the Ministry of Higher Education (MOHE) grant under the Fundamental Research Grant Scheme (FRGS) (FRGS/1/2020/SKK0/UM/03/2).

Data availability

The data are available upon reasonable request from the corresponding author.

Declarations

Competing interests

The authors declare no competing interests.

Ethics approval and consent for participation

This study was approved by the University of Malaya Medical Centre-Medical Research Ethics Committee (UMMC-MREC) (Ethics No: 20211029-10716) for patients’ retrospective clinical data collection and tissue samples acquisition. The information sheet describing the project was provided, and informed consent was obtained from all patients/donors. All procedures involving human tissue samples were carried out according to the ethical standards and institutional guidelines, and all patients’ data were anonymized prior to publication to protect individual confidentiality. Appropriate measures were taken throughout the study, ensuring full compliance with the ethical principles of the Declaration of Helsinki for research involving human participants. The use of the murine model in this in vivo study was approved by the Institutional Animal Care and Use Committee (IACUC), University of Malaya (Ethics No: 2020-231203/ORTHO/R/KG). All animal experimental procedures involved complied with the institutional guidelines and ARRIVE guidelines.