Chlamydia are a group of gram-negative, obligate intracellular pathogens, and their broad host ranges from single-celled eukaryotes to cattle, sheep and humans (1). The main species capable of commonly infecting humans include Chlamydia trachomatis, Chlamydia pneumoniae, and Chlamydia psittaci. Among them, C. trachomatis has the greatest impact on human health. The eye infection of C. trachomatis can cause trachoma, which is the main cause of preventable blindness in developing countries of the world (2). According to the data of the World Health Organization (WHO) in June 2022, at least 125 million people are living in trachoma-endemic areas face the risk of trachoma blindness (2). Besides, genitourinary infection caused by C. trachomatis may lead to venereal lymphogranuloma or infertility (3).

C. trachomatis has a unique biphasic developmental cycle that alternates between two morphologically and functionally distinct developmental stages: the small, structurally stable, infectious elementary body (EB) and the large, metabolically vigorous, replicative reticulate body (RB) (1, 4). Infection begins with attachment and internalization of EBs to host cells. RBs replicate within a membrane bound compartment - the inclusion, early genes are transcribed and EBs differentiate into RBs (~6–8 hours post-infection) (1, 5, 6). Next, effectors that mediate nutrient acquisition and maintain the viability of the host cell are expressed. The bacteria divide by binary fission and the inclusion substantially expands (~8–16 hours post-infection) (6, 7). Last, RBs gradually re-differentiate back into EBs. By host cell lysis or by extrusion of intact inclusions, EBs are released to infect neighboring host cells at the end of the developmental cycle (~24–72 hours post-infection) (1, 8, 9).

Although the sequence of the first C. trachomatis genome was published in 1998 (10), the functional analysis of proteins of C. trachomatis has been limited by the lack of tools of genetic manipulation for a long time. A major barrier to the development of genetic manipulation is its dependence on a host replication and the unique biphasic developmental cycle. RB is only present in the host cell, and it is difficult for exogenous DNA to reach the bacterial cytoplasm through the four-layers of biofilm (11). Even EB can exist in the environment, it has a hard cell wall and low metabolic activity, so it is unlikely to reabsorb and integrate foreign DNA (11).

Fortunately, in the past decade, the development and application of several tools of genetic manipulation of C. trachomatis has made some progress, greatly expanding the current research on the biological characteristics of C. trachomatis and the function analysis of its virulence factors. The first step in almost all methods of genetic manipulation of Chlamydia is transformation. Four transformation methods have been reported: electroporation, chemically induced mutagenesis, polyamidoamine dendrimers (PAMAM dendrimers), and CaCl₂ transformation (4, 12, 13). In this review, we categorize and summarize the tools of genetic manipulation that have been developed for C. trachomatis according to the methods of plasmid transformation ( Figure 1 ; Table 1 ).

Electroporation is universally effective in introducing heterologous DNA into obligate intracellular bacteria via brief electric pulses, which induce transient and reversible cell membrane permeabilization (47), and time constant and exponential decay pulse types, episomal DNA maintenance are the key features. Tam et al. successfully introduced the shuttle plasmin pPBW100 into C. trachomatis EBs by electroporation for the first time in 1994, and used it to infect McCoy cells and detected Chlamydia containing the chloramphenicol resistance gene in cell cultures (14). A similar electroporation method was used to mobilize an engineered vector into C. psittaci 6BC EBs and a efficiency was obtained with 10 µg of circular vector (1.9 ± 1.1×10⁶, n=7) (48). For the successfully transformation, EBs must be highly purified and obtained by centrifugation through Renografin density gradients, and this complex procedure maybe limit the wide adoption of electroporation by other labs. Although the optimal transformation conditions for C. trachomatis electroporation have not been fully grasped, the study of Tam shows that exogenous DNA can be introduced into EBs by electroporation and the expression of heterologous screening tags can be achieve, which laid the foundation for the development of C. trachomatis electroporation technology in the future (14).

Chemical mutagenesis is a technology that uses chemical mutagens such as base analogues, deamination agents, and alkylating agents to mutate DNA (49, 50).

C. trachomatis was subjected to low-level ethyl methanesulfonate (EMS) mutagenesis to generate chlamydiae that contained less than one mutation per genome in 2011 and a tryptophan synthase gene (trpB) null mutant incapable of avoiding the anti-microbial effect of IFN-γ–induced tryptophan starvation was isolated (15). Then mutagenesis in Chlamydia was performed by exposure of infected cells to either of the DNA alkylating compounds EMS or N-ethyl-N-nitrosourea (ENU), followed by plaque isolation of clonal strains in 2015 (4). In the study, Kokes et al. used ethyl methyl sulfonate (EMS) and N-ethyl-N-nitrosourea (ENU) to perform chemical mutagenesis on C. trachomatis to generate a mutant library, screening of mutants impaired in F-actin assembly and identifying InaC as an inclusion body membrane protein that binds host ARF and 14-3-3 proteins and regulates F-actin recombination and Golgi reorganization around vesicles (16).

Although studies have shown that it is possible to achieve either one mutation (15) or multiple mutations per genome of C. trachomatis (16–18) by adjusting different concentrations of mutagen, identifying and linking genotype and phenotype without a molecular signature is laborious and tedious, and most of the mutations identified are non-essential genes (51). Therefore, with the continuous development of genetic manipulation, the application of chemical mutagenesis will be gradually phased out.

Polyamidoamine (PAMAM) dendrimers are hyperbranched polymers with low cytotoxicity. It can not only deliver small molecules to specific sites, but also effectively transfuse biological macromolecules such as oligonucleotides and plasmid DNA into cells (52). In addition, these dendrimers can be localized in Chlamydia inclusion bodies in Chlamydia-infected cells (53), indicating that PAMAM can directly transform exogenous DNA into RBs in infected monolayers (19, 52).

In 2012, Mishra et al. used PAMAM to transfer an antisense oligonucleotide into C. trachomatis and efficiently and specifically knockdown the transcription level of the target gene (19). Then in 2013, Kannan successfully used PAMAM to transfer a plasmid (pMW82: pL2-01-pL2-01P-GFP) into C. trachomatis and successfully detected green fluorescence in the initial transformed culture (20, 48). Despite plasmid replication and GFP expression being detected within the first infection cycle which indicating a high transformation efficiency, the dendrimer-based transformation method of Chlamydia has not been broadly adopted.

Since the first utilization of CaCl₂ transformation method to stably transform the shuttle plasmid pBR325::L2 into C. trachomatis EBs by Wang et al. in 2011 (54). In the protocol, EBs were firstly incubated with plasmid DNA in CaCl₂ buffer for 30 min at room temperature and then host cells resuspended in CaCl₂ buffer were added, followed by an additional incubation for 20 min at room temperature. Due to its advantages of simple, rapid, cheap and repeatable operation, the CaCl₂ transformation method is widely used as a general transformation method for Chlamydia. For the CaCl₂ transformation, crude preparations of Chlamydia from host cell lysates exhibit more efficient than gradient purified EB preparations (54–56). Based on this method, group II intron-based targeted gene knockout (21–30), fluorescence-reported allelic exchange mutagenesis (FRAEM) (31–38), CRISPR interference (CRISPRi) (39–43) and transposon insertion mutagenesis (44–46) have been realized in C. trachomatis.

The genetic intractability of C. trachomatis has severely limited molecular dissection of virulence factors associated with intracellular parasitism and pathogenic mechanisms that promote trachoma, venereal lymphogranuloma or infertility, because there was no methods for C. trachomatis virulence determinants inactivation and/or complementation. Great progress has been made in the development of genetic manipulation of Chlamydia in the past decade, and the application of the tools of genetic manipulation has significantly impeded progress in understanding the genetic basis of the pathogen’s unique lifestyle and virulence. Moreover, the increasing genetic tractability of C. trachomatis will enable the development of new pathogen countermeasures, such as rationally designed attenuated or subunit vaccines. But some problems still remain:

Additional advances in genetic manipulation will be necessary to render Chlamydia significantly more genetically tractable. Ideally, a cell-free medium like C. burnetii (77, 78) for Chlamydia cultivation would address some issues. Omsland et al. developed a stage-specific metabolic and transcriptional activity of C. trachomatis in an axenic medium in 2012 (79), and host-free cultivation of Chlamydia may be achievable in the future. Improved electroporation conditions may be another avenue if the decreased chlamydial viability could be addressed. Overall, more rapid and definitive progress can be expected for this important and interesting intracellular parasite.

The manuscript was drafted by WW and DannL, and edited by JJ and DanL. All authors contributed to the article and approved the submitted version.