Antimicrobial Susceptibility Testing in Chlamydia trachomatis: The Current State of Evidence and a Call for More National Surveillance Studies

Abstract

Chlamydia trachomatis (C. trachomatis) remains the most common bacterial sexually transmitted agent worldwide. Although current treatment regimens are highly effective, sporadic reports of reduced antimicrobial susceptibility and treatment failure raises concerns, especially in the context of increasing global antibiotic consumption and the well-documented rise of antimicrobial resistance (AMR) in other sexually transmitted pathogens. A key factor contributing to the continued efficacy of antimicrobials against C. trachomatis is the unique biology of this species, including its obligate intracellular life cycle, reduced genome, and capacity to enter a persistent state. However, this same biology poses significant challenges to antimicrobial susceptibility testing (AST). Different national surveillance studies have consistently demonstrated low rates of resistance, confirming that C. trachomatis remains largely susceptible to first-line therapies. Nonetheless, these efforts are scarce and have also revealed significant variability in testing protocols, limited geographic coverage, as well as a lack of continuous monitoring. Since antibiotic consumption patterns differ between regions, systematic surveillance will become indispensable to detect emerging resistance trends before they translate into widespread clinical failure. This narrative review synthesizes on a molecular basis the current evidence of C. trachomatis resistance and available AST methods, evaluates findings from different national surveillance studies, and underscores the need for standardized, well-funded surveillance strategies to preserve the long-term efficacy of treatment options for chlamydiosis.

1. Introduction

Chlamydia trachomatis (C. trachomatis) is the most common sexually transmitted bacterial agent today; in 2020, 128.5 million new cases of C. trachomatis infection in adults aged 15–49 years compared to 82.4 million of new gonorrhea infections globally were estimated by World Health Organization (WHO) [1]. A recent meta-analysis involving 24 countries reported a pooled prevalence of 2.9% in the general population, with females showing a higher C. trachomatis prevalence (3.1%) compared to males (2.6%) [2]. It should be emphasized that in high-risk groups—such as female sex workers or men who have sex with men—the prevalence is significantly higher and can reach up to 30.3% [3]. Age-wise, the highest prevalence is consistently observed among individuals aged 15 to 24 years [4]. The infection is often asymptomatic, particularly in women, and if left undetected and untreated, it can lead to serious complications that may affect the reproductive health of both women and men [5,6,7]. In contrast to gonorrhea, chlamydia infection is still effectively treated with first-line antibiotics and thus far is not a concern regarding antimicrobial resistance. However, although extremely rare, isolates of C. trachomatis showing clinical and/or laboratory-observed resistance to common drugs used for treatment (i.e., tetracyclines, macrolides, and fluoroquinolones) have been described [8,9,10,11]. Moreover, due to the observed decrease in the effectiveness of the previously recommended therapy with 1 g azithromycin, some countries have already changed their guidelines for the treatment of chlamydial infection [12,13]. Therefore, in an era of growing antimicrobial resistance and increasing global antibiotic consumption, possibility for C. trachomatis resistance should not be overlooked but rather periodically monitored to ensure we are not caught unprepared. The purpose of this review is to discuss the biological characteristics underlying the unusual susceptibility of this bacterium as well as the molecular mechanisms of its resistance and to provide an overview of research on C. trachomatis susceptibility conducted to date—primarily larger, nation-wide studies. Furthermore, we aim to highlight the need for future surveillance on a regional/national level and laboratory investigations on C. trachomatis sensitivity to antimicrobials to detect and support timely reaction to possible changes in its antimicrobial sensitivity profile.

2. Biology and Molecular Basis of Resistance

C. trachomatis is an obligate intracellular bacterium that occurs in two forms: the elementary body (ET), an infectious form that can survive outside the host cell, and the reticulate body (RT), the replicative form found exclusively within the cell [14]. These two forms differ in size, structure, and type of metabolism, ultimately leading to distinct functions [15]. Transformation from one form to the other is a characteristic of the unique developmental cycle of chlamydia. ETs are 2.5 times smaller than RBs and resistant to osmotic and mechanical challenges, reflecting a specially constructed cell wall reinforced by substantial cross-linking disulfide bonds between and within outer membrane proteins [16,17,18]. Within the ET, there is a highly condensed nucleoid as a result of complete shut-down of transcription activities [15]. Upon entering the cell, genes regulating the transcription factors are activated, leading to the conversion of the ET to the RT. The intracellular phase of the chlamydial developmental cycle takes place within a specialized structure called an inclusion, which is enclosed by a membrane and provides favorable conditions for the development and division of sensitive RTs [19,20]. Another important characteristic of the RT is the containment of peptidoglycan within its cell wall [21]—an important building block of many bacteria as well as a pertinent target site for antimicrobial drugs. More precisely, although it was previously believed that Chlamydia does not contain peptidoglycan, recent studies have visualized it in the septum during the binary division of the RT [22] or confirmed its presence by analyzing peptides obtained from lysates of cells infected with C. trachomatis [23]. It is well known that under certain stressful conditions, such as tryptophan or iron starvation, exposure to interferon gamma, exposure to penicillin, and even concurrent herpes virus infection, the RB transforms into an enlarged aberrant body (AB) and enters a persistent state [24,25,26]. This is a reversible process, and after the stress stimulus is removed, the AB, which does not replicate but is still viable, exits the persistent state and assumes the form and characteristics of a normal RB [24]. Even more intriguing is the discovery that azithromycin, one of the first drugs of choice for chlamydial treatment [27], induces the appearance of aberrant bodies in an in vitro model and entry into a persistent state, which is dependent upon dose and post-infection time [28]. Moreover, in vitro-induced persistent infection by azithromycin was found to be not related to typical mutation for macrolide resistance mutations on 23S rRNA, L4, or L22 protein genes, and it was completely recoverable after removal of the drug. Since the reduced susceptibility of clinical isolates of C. trachomatis observed under laboratory conditions is not always result of mutations in specific genes [9], the possibility of multiple resistance mechanisms arises, some of which may be closely linked to the bacterium’s ability to enter a persistent state. In any case, the role of the persistent state in the pathogenesis of chronic infection, recurrence, as well as antimicrobial resistance remains to be elucidated.

Like other members of the Chlamydiaceae family, and akin to the other obligate intracellular bacteria, C. trachomatis has a reduced genome that is a consequence of genome streamlining during co-evolution within their eukaryotic hosts [29,30,31]. It possesses a circularized chromosome of approximately 1.05 Mbp and a highly conserved plasmid of approximately 7.5 kbp in size [31,32], the latter being the key virulence factor and determinant of pathogenicity of chlamydia species. Plasmid-less strains of C. trachomatis are rare but have been observed, with a consequent reduction in virulence and pathogenicity [33,34]. Whole-genome sequencing has significantly advanced our understanding of the organization and composition of chlamydia genomes, including C. trachomatis [31,35]. However, it has contributed little to our knowledge of chlamydial antibiotic resistance genes.

Previous studies focused on antimicrobial resistance (AMR) genes and their mechanisms in C. trachomatis have identified several regions where mutations have been detected in strains with reduced susceptibility [35,36]. Regions and genes related to AMR for a particular group of antibiotics as well as pertinent studies are listed in

Table 1. Specific genes and genome region mutated in antibiotic resistant strains of C. trachomatis.

Antimicrobial Drug	Genes and Regions with Detected Mutations	References
Macrolides	23S rRNA, rpID L4 protein gene, rpIV L22 protein gene	Misyurina 2004 [37], Binet 2007 [41], Zhu 2010 [42], Yiang 2015 [43], Deguchi 2018 [44]
Tetracyclines	ompB/porB gene, tet(M) gene for ribosomal protection protein	O’Neill 2013 [45], Li 2017 [46], Shao 2020 [47]
Fluoroquionolones	gyrA quinolone-resistance-determining region (QRDR), ParC gene for topoisomerase IV subunit C, ygeD gene for efflux protein	Dessus-Babus 1998 [48], Morrissey 2002 [49], Yokoi 2004 [50], Misiurina 2004 [51], Deguchi 2018 [44]
Rifampin	rpoB gene coding beta-subunit of DNA-dependent RNA polymerase	Dreses-Werringloer 2003 [52], Kutlin 2005 [53], Suchland 2005 [54], Rupp 2008 [55]

. However, as previously mentioned, it should be noted that strains with reduced susceptibility to antibiotics have also been detected despite the absence of detectable mutations in their genome [11,37]. Moreover, the only chlamydial species to have naturally obtained a resistance gene integrated as a genomic island (e.g., tetC encoding for efflux pump) is C. suis [38,39]. Although in vitro co-culture studies have demonstrated that horizontal gene transfer can occur between species (e.g., C. suis and C. trachomatis) [40], these observations have not yet been found in clinical isolates [31].

3. Methods for C. trachomatis Antimicrobial Susceptibility Testing

Antimicrobial susceptibility testing for C. trachomatis deviates from standard bacteriology, requiring assessment of its replication within cells via detection of intracellular inclusions [56,57,58]. Here, cell culture systems are used, predominantly with McCoy cells, although HeLa, HEp-2, and HL (and less commonly BGMK or Vero cells) are also used [56,57]. Immunofluorescence staining identifies these inclusions, with optimal yields influenced by factors like high glucose, neutral pH, as well as elevated centrifugation temperatures (33–35 °C). Variables such as inoculum size and timing of antibiotic administration and removal alongside laboratory conditions like temperature and pH may affect results, though their impact is underexplored; however, the quality control of cell lines is essential [57]. Unlike natural infections where antibiotics act post infection and inflammation, in vitro testing applies drugs concurrently or soon after infection, potentially missing viable but non-infectious states.

Determining minimal inhibitory concentrations (or MICs) in such cell cultures is complicated by subjective interpretation of inclusions, with MIC_TP (transition point MIC) serving as a marker for the concentration at which 90% or more of the inclusions exhibit abnormal morphology [56,58]. This process can be further enhanced by employing a secondary passage of the culture onto fresh cells without the presence of the antimicrobial, aiming to ascertain the minimal chlamydicidal concentration (MCC), indicative of complete organism eradication. Such culture-based approaches are instrumental not only in quantifying the count of infectious EBs, known as recoverable inclusion-forming units (IFU), but also in evaluating the antimicrobial’s efficacy in preventing the formation of inclusions. Still, it has to be noted that the interpretation of elevated MICs in C. trachomatis is very cumbersome without breakpoints or MIC distribution data for wild-type strains and those after treatment failure [56]. And although there were early attempts to standardize the process, the approach can vary substantially between different laboratories [59].

Alongside this classic approach in McCoy cells and MIC determination, a research team from China recently introduced the fractional inhibitory concentration (FIC) method, using a checkerboard assay [60]. This was pursued to evaluate the interaction of antibiotic combinations (i.e., azithromycin plus moxifloxacin, azithromycin plus minocycline, or minocycline plus moxifloxacin) against C. trachomatis by serially diluting one antibiotic along the ordinate and the other along the abscissa of 96-well plates, overlaying inoculated wells with both drugs in two-fold dilutions. In their approach, FIC was calculated as the sum of FIC A and FIC B, where the latter two can be seen as the MICs of each drug in combination divided by their MICs alone, with synergy defined as FIC ≤ 0.5, indifference as 0.5 < FIC < 2, and antagonism as FIC ≥ 2 [60].

However, evidence points to the fact that identifying resistance genes may better account for the high treatment failure rates in urogenital C. trachomatis infections compared to MIC results, emphasizing the importance of genetic testing for antimicrobial resistance in affected patients [47]. This is why, for some time now, there has been an emphasis on molecular methods for the purposes of C. trachomatis susceptibility testing. And one of the pioneering studies in using molecular approaches for antimicrobial susceptibility testing of C. trachomatis was the work performed by Cross and co-authors [61], who introduced and evaluated a reverse transcriptase–PCR (RT-PCR)-based method for comparing it to the conventional method of cell culture and immunofluorescence (IF). The RT-PCR method aimed to improve sensitivity and reduce subjectivity in determining MICs. C. trachomatis still has to be cultured in McCoy cells with varying concentrations of antimicrobials, and after 48 h, RNA can be extracted, so RT-PCR targeting the DnaK gene can be performed [61]. The MIC is then determined as the lowest concentration of antibiotic that inhibited the appearance of a PCR product. Cross et al. showed that the RT-PCR method consistently yielded higher MIC values compared to IF, indicating greater sensitivity, and suggested that RT-PCR detects viable Chlamydiae at lower levels of replication than IF [61]. Furthermore, it may even detect aberrant forms of Chlamydiae that IF misses. Hence, it can be seen as a more sensitive and objective method for assessing C. trachomatis susceptibility, particularly for detecting low-level replication and potential persistence.

The application of PCR methods extends beyond mere detection, encompassing the direct quantification of C. trachomatis from clinical specimens and offering a more nuanced understanding of the bacterial load. This is exemplified in the work of a Hungarian research group that pioneered a direct qPCR-based methodology [58,62]. Their innovative approach bypasses the conventional nucleic acid purification step, instead opting for a straightforward lysis of infected host cells through freeze–thaw cycles, followed by the direct application of EvaGreen-based qPCR on these lysates. This method has proven effective in assessing the impact of various antibiotics, antiseptics, and growth-promoting compounds on Chlamydia, showcasing its versatility [58,62].

A multicentric research team from the Netherlands, U.S., and India developed a specific 23S rRNA gene PCR assay to detect known macrolide resistance-associated mutations (RAMs) directly from urogenital and rectal samples, bypassing the need for prior culture and thus offering a high-throughput, less technically demanding alternative to C. trachomatis culture for genotypic resistance surveillance [63]. The assay’s suitability for clinical samples was validated through systematic in silico analysis and specificity testing, showing no false positives in 40 C. trachomatis-negative swabs (20 vaginal and 20 rectal) or with a specificity panel of bacteria except for C. suis and Chlamydia muridarum; however, clinical sample testing confirmed detection was specific to C. trachomatis sequences [63]. With an analytical sensitivity consistently detecting the C. trachomatis 23S rRNA gene at 10³ IFU/mL, the assay demonstrates strong performance for direct susceptibility screening.

Very recently, a team from Portugal used a multiplex NGS-based approach, focusing on detecting resistance markers in the 23S rRNA (mutations at A2057, A2058, and A2059 for macrolide resistance) and gyrA/parC genes (QRDR substitutions for fluoroquinolone resistance) [64]. The amplification is carried out via nested and multiplex PCR and further sequenced with Sanger and Illumina NextSeq 550 platforms before tools like INSaFLU and MEGA11 can be used against reference strains. The benefit of their approach is also the possibility to survey genomic diversity through pmpH, CT105, and CT442 loci, identifying strain-specific insertions and phylogenetic relationships and in turn providing very detailed insights into resistance mechanisms and genomic backgrounds of C. trachomatis [64].

It is already evident that, in the future, whole-genome sequencing may offer insights into C. trachomatis strain evolution and the pertaining AMR patterns. Büttner et al. [65] evaluated four methods using positive clinical samples and showed how target enrichment with custom SureSelect probes targeting C. trachomatis, followed by Illumina sequencing, can successfully produce whole genomes from 64% of positive samples. By enabling the analysis of circulating strains and genotypic resistance, it may indeed be a promising tool for susceptibility testing, surveillance, and subsequent public health strategies.

There are also some alternative approaches, but their use has not fully materialized in quotidian practice. For example, flow cytometry was considered a promising alternative for C. trachomatis susceptibility testing by providing an objective, reproducible measure of antibiotic efficacy, potentially enhancing surveillance efforts in the face of evolving resistance concerns [66]. Unlike traditional microscopy, it quantifies chlamydial inhibition through fluorescence intensity, reducing subjectivity and improving precision in assessing drug activity. Dessus-Babus and co-authors [66] compared flow cytometry with the standard microscopic MIC method, testing the L2 reference strain and 13 clinical isolates from six patients with recurrent infections in McCoy cells infected with 10⁵ IFU/mL and treated with doxycycline, ofloxacin, and erythromycin dilutions. Here, flow cytometry measured the mean fluorescence intensity (MFI) of stained inclusions, yielding equivalent (but imprecise) MIC endpoints to microscopy. However, the inhibitory concentration 50 (IC50)—the dose reducing MFI by 50%—offered a more accurate, reproducible metric, varying by one to three dilutions from MICs. Nonetheless, while specific and objective, flow cytometry’s lower sensitivity misses low-level “heterotypic resistance”, and its costly equipment and time-intensive processing limit widespread use, which may be reasons why it was never as popular for use as cell culture or molecular methods.

4. C. trachomatis Sensitivity Surveillance Efforts

Surveillance of C. trachomatis susceptibility across different countries should be seen as a pivotal step even though clinically relevant resistance is debated [56,67], as it provides important insights into potential shifts in antimicrobial efficacy. There is a variability in local antibiotic use, strain diversity, and diagnostic practices that actually necessitates country-specific monitoring to detect emerging resistance patterns that might not yet manifest as widespread clinical failure but could signal future challenges—particularly in high-burden regions and regions with a high number of antibiotic prescriptions. In our review, only surveillance studies with more than 20 specimens were included to ensure robust sample sizes, enabling meaningful comparisons across diverse populations and healthcare systems.

In what is considered one of the earliest examples of larger surveillance efforts of C. trachomatis antimicrobial sensitivity in the literature, researchers from Israel evaluated the in vitro efficacy of clarithromycin, azithromycin, roxithromycin, erythromycin, doxycycline, and tetracycline against 50 C. trachomatis clinical isolates using a cycloheximide-treated McCoy cell tissue culture system to determine MICs and minimal bactericidal concentrations (MBCs) [68]. All isolates showed MICs of ≤0.015 μg/mL for clarithromycin, ≤0.125 μg/mL for roxithromycin and azithromycin, and ≤0.25 μg/mL for erythromycin and doxycycline, while 44% required 0.5 μg/mL of tetracycline for inhibition [68]. MBCs reached up to 4 μg/mL for doxycycline (8% of isolates) and tetracycline (4%), but azithromycin, a CDC-recommended treatment, demonstrated significantly superior in vitro activity compared to erythromycin and doxycycline.

Another laudable and very broad surveillance effort on C. trachomatis sensitivity was conducted in Japan [69]. In this study, urethral discharge specimens were collected from patients with urethritis in 51 hospitals and clinics in 2009 and 38 in 2012. MIC was determined for 19 isolates in 2009 and 39 in 2012. In both years, the MICs (MIC₉₀) for several antibiotics were as follows: ciprofloxacin (2 μg/mL in 2009; 1 μg/mL in 2012), levofloxacin (0.5 μg/mL), tosufloxacin (0.125 μg/mL), sitafloxacin (0.063 μg/mL), doxycycline (0.125 μg/mL), minocycline (0.125 μg/mL), erythromycin (0.016 μg/mL), clarithromycin (0.063 μg/mL), and azithromycin (0.063 μg/mL) [69]. In this study, there were no resistant strains against macrolide and/or tetracycline agents in Japan.

This was followed more recently by the second broad nationwide surveillance of the antimicrobial susceptibility of C. trachomatis in Japan from 26 hospitals and clinics in 2016 and 2017 [70], which (once again) did not identify any strains resistant to aforementioned antimicrobial agents. MIC values were comparable with the previous research endeavor [70]. Solithromycin was introduced into this study and showed the lowest MIC (0.016 μg/mL) in comparison to other antimicrobials tested; however, azithromycin’s MIC was slightly higher than in the first surveillance, though possibly within acceptable error margins, warranting the need for ongoing monitoring.

As part of a national surveillance effort in Croatia—a country with Europe’s highest azithromycin consumption and elevated antibiotic prescription rates—Ljubin-Sternak et al. [71] initially evaluated the in vitro susceptibility of 24 urogenital Chlamydia trachomatis strains to azithromycin and doxycycline, the recommended first-line treatments. Conducted using a McCoy cell culture system, they tested 14 cervical, 9 urethral, and 1 prostatic secretion isolate, finding all strains susceptible, with MICs of 0.064–0.125 μg/mL for azithromycin and 0.016–0.064 μg/mL for doxycycline and MCCs of 0.064–2.0 μg/mL and 0.032–1.0 μg/mL, respectively [71]. Despite reported clinical treatment failures linked to resistance elsewhere, no resistance was detected, underscoring the importance of ongoing surveillance to monitor potential shifts in antimicrobial susceptibility amid high antibiotic use.

This was broadened several years later by Meštrović et al. [72], with a total of 40 tested C. trachomatis strains from Croatia showing susceptibility to the antibiotics evaluated (MIC < 4 μg/mL), with no evidence of homotypic or heterotypic resistance patterns. The MCCs were either equivalent to or 1–5 dilutions greater than the MICs. In vitro testing demonstrated statistically significant variations in the efficacy of the antimicrobial agents in laboratory conditions [72]. A notable correlation was observed between MCCs for two pairs of antibiotics: azithromycin with levofloxacin and doxycycline with levofloxacin. When comparing median values across different clinical sample types, no significant differences emerged. And although not statistically significant, a trend toward higher MICs for azithromycin was observed, suggesting the need for further investigation [72].

In two surveillance studies from Tanzania, researchers investigated potential resistance development in ocular C. trachomatis infections following three annual mass drug administrations (MDA). In an earlier study from Kahe Mpya, Solomon et al. [73] assessed C. trachomatis susceptibility to azithromycin and tetracycline before and two months after mass antibiotic treatment. All chlamydial isolates had MCCs of azithromycin ≤ 2.0 μg/mL, well below the mean conjunctival tissue concentration of 24.5 (±9.7) μg/g measured in four patients 24 h after a 1 g dose, indicating sufficient drug levels [73]. Pre-treatment MCCs for tetracycline were ≤0.5 μg/mL, dropping to ≤0.06 μg/mL post treatment. Hence, mass antibiotic treatment did not induce clinically or programmatically significant changes in C. trachomatis susceptibility to either azithromycin or tetracycline in this community. In a study by West et al. [74] in 32 endemic communities, one year post MDA, there were 107 children infected; after another MDA, 90 were re-evaluated at two months, with 30 still infected. Paired isolates from 15 children before and after MDA showed no significant resistance, with average MICs of 0.26 µg/mL and 0.20 µg/mL for azithromycin, respectively, and all MICs ≤ 0.50 µg/mL [74]. Of note, infectious loads were similar between consistently infected children and those whose infections cleared, indicating that persistent infections were not due to azithromycin resistance or higher pre-treatment bacterial loads.

In a surveillance study from Italy, Foschi et al. [75] evaluated the in vitro antimicrobial susceptibility of 20 C. trachomatis strains—representing common genovars D, E, F, and G—in HeLa (endocervical) and Caco-2 (colorectal) cell lines to mimic urogenital and rectal infection sites. Susceptibility to azithromycin, erythromycin, doxycycline, and levofloxacin was tested, also exploring correlations between MIC values and bacterial load. Results showed that macrolides (azithromycin and erythromycin) had higher MIC and MBC values (by 2-fold dilutions) in Caco-2 cells compared to HeLa cells, while doxycycline and levofloxacin exhibited no notable differences between cell lines; additionally, azithromycin MICs in Caco-2 cells rose significantly with higher C. trachomatis elementary body loads [75]. Their findings suggest that elevated azithromycin MICs in colorectal cells, combined with load-dependent increases, may contribute to treatment failures for rectal C. trachomatis infections.

A multicenter study that involved seven Spanish National Health Service hospitals aimed to assess the phenotypic and genotypic susceptibility of C. trachomatis to key antimicrobials (macrolides, doxycycline, and quinolones) in isolates from patients around the country with treatment failure, excluding reinfection [76]. From 2018 to 2019, 73 isolates were collected. Phenotypic testing was conducted on 69 specimens using McCoy cell monolayers, while genotypic analysis involved sequencing the 23S rRNA gene (positions 2057, 2058, 2059, and 2611) for macrolide resistance, the gyrA gene (G248T mutation) for quinolone resistance, and RT-PCR for the tet(C) gene for tetracycline resistance [76]. Phenotypic testing succeeded for 10 isolates, all sensitive (MICs ≤ 0.125 mg/L for azithromycin; ≤0.064 mg/L for doxycycline). Among 66 sequenced isolates, no resistance mutations were detected in 23S rRNA, gyrA, or tet(C). More specifically, no evidence of genomic resistance was found.

A very recent national surveillance study by the Portuguese National Reference Laboratory for STIs examined 502 C. trachomatis-positive samples, primarily anorectal exudates, to assess genetic markers of AMR to macrolides and fluoroquinolones, with the use of multiplex PCR and next-generation sequencing (NGS) [64]. No samples showed 23S rRNA mutations linked to macrolide resistance, but three (0.6%) had a GyrA Ser83Ile mutation potentially conferring fluoroquinolone resistance. The study also successfully differentiated LGV from non-LGV strains and identified L2/L2b genomic backbones, with a notable insertion upstream of CT105 showing promise as a rapid LGV diagnostic target [64]. Overall, it suggests low dissemination of AMR genetic determinants in clinical C. trachomatis strains in Portugal.

5. Bridging Concept and Practice in the Implementation of National Surveillance Studies

Despite the current consensus in the literature that C. trachomatis remains largely susceptible to first-line antimicrobials, the global landscape of AMR would benefit from a proactive surveillance in this case as well to prevent the emergence of undetected resistance patterns. The lack of standardized and routine antimicrobial sensitivity testing for C. trachomatis contributes to a gap in knowledge regarding potential shifts in its resistance profile [58], most notably in high-prevalence regions with extensive antibiotic usage. Given the documented cases of reduced susceptibility and occasional treatment failures [8,9,10,11], it can be argued that there is a need for much more national surveillance studies that systematically assess C. trachomatis sensitivity to commonly prescribed antimicrobial agents. A proposed surveillance algorithm to conduct such monitoring is presented in

Table 2. Proposed surveillance algorithm to monitor Chlamydia trachomatis (C. trachomatis) antimicrobial susceptibility trends.

Step	Process	Description
1	Clinical case identification or screening programs	Recognizing suspected C. trachomatis infections in primary, secondary, and tertiary care as well as in STI clinics in symptomatic individuals; conducting targeted screening programs to detect C. trachomatis infections in asymptomatic individuals
2	Sample collection and transport	Collecting urogenital, anorectal, and pharyngeal specimen for testing while ensuring proper storage and transport to clinical microbiology laboratories
3	Laboratory confirmation of infection	Performing nucleic acid amplification tests for diagnosis; if positive, alerting clinicians in step 1 for more samples (if necessary) to conduct AST
4	Conducting AST of isolates	Conducting culture-based AST and/or molecular assays to detect resistance markers (contingent upon availability in the reference laboratory)
5	Data integration and analysis	Aggregating results to identify shifts in MICs and other resistance patterns/trends across different settings; developing regional and national antibiograms
6	Genomic investigation	Employing whole-genome sequencing and next-generation sequencing to monitor resistance-associated mutations and strain evolution (contingent upon availability)
7	Surveillance reporting	Sharing findings with healthcare providers on all levels, public health agencies, and relevant stakeholders to inform further decisions
8	Guideline updates and policy	Adjusting treatment protocols (if necessary) and national healthcare policies based on the analysis of surveillance data to guide clinical action and policy development

Abbreviations: AST—antimicrobial susceptibility testing; MIC—minimal inhibitory concentration; STI—sexually transmitted infections.

and a simplified diagram in Figure 1.

A viable strategy should integrate both phenotypic and genotypic approaches to obtain a full picture of antimicrobial susceptibility trends. Traditional cell culture-based AST methods remain valuable for assessing MICs and MCCs, yet their labor-intensive nature and lack of standardized breakpoints hinder their routine application [56,57,70]. On the other hand, molecular methods—with examples such as PCR-based resistance marker detection and next-generation sequencing (NGS)—offer rapid, high-throughput insights into genetic determinants of resistance [31,64,65]. Combining these methodologies in a multipronged approach, where culture-based methods validate molecular findings, may be one potential framework for surveillance.

The variability in antibiotic prescribing practices [77,78] means that regional and/or national surveillance programs should prioritize the inclusion of diverse clinical isolates from symptomatic and asymptomatic infections as well as from both urogenital and extragenital sites. This is especially important in regions with high azithromycin or tetracycline consumption, where selective pressure may drive the emergence of resistance mechanisms that have yet to be fully characterized [68,72]. Additionally, in the future, periodic reassessment of treatment guidelines may be more readily informed by MICs and MCCs rather than by historical efficacy alone.

Collaboration among microbiology laboratories, public health agencies, and clinical practitioners is essential for establishing a sustainable and effective surveillance network for C. trachomatis antimicrobial susceptibility [79]. Considering all the challenges associated with C. trachomatis culture-based AST, which is not routinely performed in clinical microbiology laboratories, there is a need to include institutes for public health and national/regional reference laboratories (as is the case in infectious disease surveillance in general) and adequately fund them [80]. They should also take the lead in ensuring diagnostic stewardship and data quality across different geographic regions and healthcare systems [81]. Public health agencies should also be tasked with facilitating coordination between laboratories, providing logistical support, funding, and policy guidance to integrate sensitivity profiles into broader STI surveillance and control programs.

Equally significant is the involvement of clinical practitioners who manage chlamydial infections on the front lines. They should be included in surveillance efforts by contributing clinical data and recognizing potential treatment failures but also advocating for confirmatory testing in cases of persistent infections [82]. Training programs for healthcare providers, which are already part of continuing medical education to increase screening yields [83], should additionally emphasize the importance of monitoring treatment efficacy and implementing follow-up testing in high-risk populations. Additionally, collaborations with primary care providers and sexual health services (primarily STI clinics) are indispensable if we aim for the surveillance data to reflect real-world treatment outcomes.

To achieve consistency and reproducibility across different settings, standardized protocols for C. trachomatis AST must be developed and widely adopted, which is still something the medical community needs to work on. These should encompass not only traditional cell culture-based approaches but also molecular assays that detect resistance-associated mutations in key genes such as 23S rRNA (for macrolides), ompB/porB and tet(M) (for tetracyclines), gyrA/parC (for fluoroquinolones), and efflux pump-related genes (Table 1). The establishment of international reference panels for AST, including wild-type and resistant C. trachomatis strains, would allow laboratories to calibrate their methods and interpret results with greater reliability. A good step forward would be the definition epidemiological cut-off values (ECOFFs) and clinical breakpoints for C. trachomatis to establish resistance [84], as this is the way to differentiate natural MIC variability from true resistance.

Beyond traditional AST, genomic surveillance initiatives should be incorporated into routine clinical and epidemiological investigations to enable the early detection of resistance-associated mutations and their potential transmission dynamics. Albeit currently very expensive and thus unattainable by many laboratories, with time, this may become more affordable to use for this purpose. For example, WGS and targeted next-generation sequencing approaches are indeed powerful tools to identify emerging resistance patterns and track strain evolution in different populations [85]. By establishing centralized databases for C. trachomatis genomic data, researchers and public health authorities would be able monitor phylogenetic relationships, detect novel resistance determinants, and also assess how genetic variations may influence treatment outcomes. Ideally, genomic surveillance data should be integrated with clinical metadata (as effectively demonstrated for some other pathogens [86]), which include patient demographics, treatment histories, co-infections and co-morbidities, to provide a more comprehensive understanding of C. trachomatis epidemiology and antimicrobial susceptibility trends.

6. Conclusions

While C. trachomatis continues to exhibit high susceptibility to first-line antimicrobials, the growing global threat of AMR means that there is a need to stay vigilant. Evidence from various national surveillance studies (conducted in countries such as Japan, Croatia, Spain, Portugal, and Tanzania) has consistently demonstrated low rates of resistance and stable MIC values—even in regions with high antibiotic consumption. These studies provide invaluable baseline data, yet they also reveal gaps in standardized testing protocols and molecular surveillance. This is why we emphasize not only the unique biological features that complicate AST in this pathogen but also the urgent need for more structured, systematic, and regionally tailored surveillance programs to detect potential early signals of antimicrobial resistance. The call for more national surveillance studies is reflected in our strong recommendation to develop standardized, multidisciplinary approaches that combine phenotypic, molecular, and genomic methodologies in countries with high antibiotic usage or with documented treatment failures.

Consequently, to ensure sustained efficacy of existing treatment regimens, it will be essential to adopt comprehensive and standardized surveillance strategies that integrate the aforementioned techniques; however, limited funding and resource allocation as well as inadequate attention to this pathogen remain significant obstacles in achieving this goal. Coordinated efforts among clinical practitioners, microbiology laboratories, and national public health institutions is also needed for establishing robust surveillance networks capable of early detection and response if we are ever faced with clinically relevant resistance patterns. It is clear that proactive monitoring (coupled with periodic reassessment of clinical guidelines based on surveillance data) has the propensity to preserve effective treatment options and prevent potential silent emergence of AMR in C. trachomatis, which should be seen as a responsibility of each country.