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Expression and structure of the Chlamydia trachomatis DksA ortholog

Abstract Chlamydia trachomatis is a bacterial obligate intracellular parasite and a significant cause of human disease, including sexually transmitted infections and trachoma. The bacterial RNA polymerase-binding protein DksA is a transcription factor integral to the multicomponent bacterial stress response pathway known as the stringent response. The genome of C. trachomatis encodes a DksA ortholog (DksACt) that is maximally expressed at 15–20 h post infection, a time frame correlating with the onset of transition between the replicative reticulate body (RB) and infectious elementary body (EB) forms of the pathogen. Ectopic overexpression of DksACt in C. trachomatis prior to RB–EB transitions during infection of HeLa cells resulted in a 39.3% reduction in overall replication (yield) and a 49.6% reduction in recovered EBs. While the overall domain organization of DksACt is similar to the DksA ortholog of Escherichia coli (DksAEc), DksACt did not functionally complement DksAEc. Transcription of dksACt is regulated by tandem promoters, one of which also controls expression of nrdR, encoding a negative regulator of deoxyribonucleotide biosynthesis. The phenotype resulting from ectopic expression of DksACt and the correlation between dksACt and nrdR expression is consistent with a role for DksACt in the C. trachomatis developmental cycle.


Introduction
Chlamydia trachomatis is a human-adapted bacterial obligate intracellular parasite and different pathotypes (aka serovars) of the pathogen cause distinct diseases. Urogenital serovars of C. trachomatis are a major cause of sexually transmitted infections (Stamm 1999, Rowley et al. 2012, while ocular serovars can cause trachoma (Burton and Mabey 2009), a leading cause of preventable blindness most prevalent in developing areas with poor sanitation. During infection of cultured host cells, C. trachomatis transitions between two physiologically distinct cell forms: the infectious but non-replicative elementary body (EB) and the replicative but noninfectious reticulate body (RB) (Shaw et al. 2000). While transition between cell forms is critical for replication and generation of infectious progeny, little is known about the trigger(s) or regulatory mechanism(s) that influence and control chlamydial morphological differentiation. Temporally restricted expression of different classes of genes during the chlamydial developmental cycle (Shaw et al. 2000, Belland et al. 2003 and recent genetic analysis identifying several genes with apparent functions in chlamydial development (Brothwell et al. 2016) suggest that EB-RB-EB transitions can occur in response to various stimuli and that response mechanisms controlling chlamydial development are redundant.
Chlamydia trachomatis has undergone genome reduction at the expense of reduced metabolic capacity (Stephens 1998). As a result, numerous essential nutrients are scavenged from the host cell during infection to allow for pathogen replication. In addition to biosynthetic deficiencies, certain regulatory pathways appear reduced to a single component of the canonical machinery. Consequently, C. trachomatis may have evolved to use these single components or 'orphan regulators' in place of certain canonical regulatory cascades.
In several bacteria, including Escherichia coli (Magnusson et al. 2007, Vinella et al. 2012, Kessler et al. 2017, Pseudomonas aeruginosa (Perron et al. 2005) and Legionella pneumophila (Dalebroux et al. 2010), DksA is a relatively small (<160 residue) RNA polymerase (RNAP)-binding protein and global transcriptional regulator that integrates responses to diverse stimuli, including nutrient availability (Paul et al. 2005), via a multicomponent response mechanism referred to as the stringent response (Cashel and Gallant 1969, Cashel et al. 1996, Potrykus and Cashel 2008, Traxler et al. 2008). The C. trachomatis genome encodes a DksA ortholog, CTL0664 (DksA Ct ). The signaling nucleotide guanosine tetraphosphate (ppGpp) can be produced in response to nutrient (e.g. amino acid) starvation and functions with other regulatory components of the stringent response, including DksA (Magnusson et al. 2005, 2007, Ferullo and Lovett 2008, Traxler et al. 2008, Geiger et al. 2010. Chlamydia trachomatis does not encode any apparent ortholog for proteins involved with ppGpp biosynthesis or breakdown (Mittenhuber 2001, Hauryliuk et al. 2015, and attempts to detect ppGpp in C. trachomatis have been unsuccessful (data not shown). The ability of DksA to function independently of ppGpp in E. coli and Rhodobacter sphaeroides (Magnusson et al. 2007, Vinella et al. 2012, Lennon et al. 2014 demonstrates that DksA is capable of functioning as an independent regulator. Additionally, in the biphasic developmental cycle of L. pneumophila, DksA has been shown to govern bacterial differentiation from the replicative form to the virulent form in cooperation with ppGpp (Dalebroux et al. 2010). We hypothesize that dksA Ct is maintained as a gene associated with RB-EB transitions in the chlamydial developmental cycle.
To probe the role of dksA Ct in chlamydial biology, we determined the expression of DksA Ct during the chlamydial developmental cycle and tested the effect of DksA Ct expression on C. trachomatis replication and EB generation following ectopic expression of dksA Ct . Additionally, we resolved the 3D structure of DksA Ct , assessed the ability of DksA Ct to functionally complement DksA Ec and identified promoter sites for dksA Ct . To determine whether DksACt expression is temporally regulated, expression of the native protein was determined at critical stages of the developmental cycle. (A) Western blot of C. trachomatis whole-cell lysates normalized to GE. Time points were chosen to include cell populations that were predominantly RBs (15 hpi, 20 hpi), mixed RBs and EBs (30 hpi), and predominantly EBs (45 hpi) (n = 2). (B) IFU assay depicting the relative abundance of infectious EBs at different time points of the chlamydial developmental cycle (n = 3).

Native expression of DksA Ct is maximal at the intermediate-late stages of the chlamydial developmental cycle
During infection of HeLa cells, C. trachomatis L2 undergoes a predictable developmental cycle in which log phase RBs initiate transition to the EB form approximately 18 h post infection (hpi) (Grieshaber et al. 2018). To determine whether DksA Ct expression is temporally synchronized with RB-EB transitions, bacteria were isolated from infected HeLa cells at time points correlating with critical stages of the chlamydial developmental cycle and protein expression determined by western blot using a DksA Ctspecific antibody (Fig. 1A). DksA Ct expression was normalized to the number of bacteria by measuring genome equivalents (GE) (Skipp et al. 2016). While minimal (if any) DksA Ct was detected in bacteria recovered at 15 hpi, a time point correlating with replicating RBs, maximal expression was observed at 20 hpi, a time point in the developmental cycle coincident with initiation of RB to EB morphological transitions. A similar expression pattern was previously reported for DksA in C. pneumoniae (Mukhopadhyay et al. 2006), indicating that DksA expression profiles are conserved between pathogenic Chlamydia species. DksA Ct was detected at moderately reduced levels at 30 and 45 hpi. Western blot analysis of EBs purified by density gradient centrifugation after sonication confirmed expression of DksA Ct in this cell form (data not shown). The dynamic expression pattern of DksA Ct is unique compared with the stable expression observed for DksA Ec (Paul et al. 2004, Brown et al. 2002. Because the RB form contains more protein per cell than the EB, the method of normalization will influence the measured level of expression of any protein in C. trachomatis. The RB contains ∼10-fold more protein compared with the EB on a per cell basis (Omsland et al. 2012), indicating DksA Ct levels remain high in the EB form. To determine the relative population of infectious EBs in infected HeLa cultures, inclusion forming unit (IFU) assays were performed (Fig. 1B). Few detected inclusions following infection with bacteria harvested at 15-20 hpi indicate that initiation of DksA Ct expression occurs at a time point when the intracellular chlamydial population predominantly consists of replicating RBs. Development of inclusions in cultures infected with bacteria recovered at 25 hpi was used as a positive control for the assay.

Overexpression of DksA Ct prior to RB-EB transition reduces both replication and production of infectious progeny
To test whether early expression of dksA Ct affects replication and/or production of infectious EBs, C. trachomatis was transformed with the inducible riboswitch construct pBOMB4::E-Riboswitch-dksA Ct (R-dksA Ct ). Additionally, to ensure any phenotype observed upon induction of the R-dksA Ct construct is not a nonspecific effect of protein expression, HeLa cultures were infected with C. trachomatis transformed with p2TK2-SW2::E-Riboswitch-Clover (R-Clover) (Grieshaber et al, 2022) and expression of Clover, a variant for green fluorescent protein, used as a control. Replication was measured via analysis of GE in samples harvested at 30 hpi in the absence or presence of 0.5 mM theophylline, added at the time of infection. Induced expression of DksA Ct resulted in a 39.3% reduction in GE compared with non-induced control cultures ( Fig. 2A). No significant effect on replication was observed with ectopic expression of Clover. These data suggest that the negative impact of early expression of DksA Ct on replication is specific to DksA Ct activity and is not due to nonspecific effects of induced protein expression.
To assess whether early expression of dksA Ct similarly impacts the chlamydial developmental cycle, generation of infectious progeny was measured by performing IFU assays with inocula normalized to GE. Induction of R-dksA Ct resulted in a 49.6% reduction in recovered IFUs as compared with non-treated control cultures (Fig. 2B). Relative EB abundance quantified via an IFU assay for bacteria transformed with R-Clover resulted in a 13.6% reduction in recovered IFUs in the presence of inducer (Fig. 2B). To determine whether the reduction in IFUs observed following induction of R-Clover was related to the protein and/or use of theophylline, the effect of theophylline on wild-type C. trachomatis L2 was determined. Although wild-type bacteria showed statistically significant differences in both replication and recovered IFUs in response to theophylline (Fig. S1, Supporting Information), the effect was less than that observed for bacteria transformed with R-dksA Ct , consistent with an effect of DksA Ct in chlamydial replication and development (GE: 18% vs 39.3%; IFU: 17.7% vs 49.6%). Compared with induction of R-Clover, induction of R-dksA Ct resulted in a statistically significant 36% reduction in recovered IFUs (Fig. 2B). Successful induction of R-dksA Ct and R-Clover by theophylline was confirmed via western blot or fluorescence microscopy, respectively (Fig. S2, Supporting Information). reduction in GE when compared with the non-induced R-dksACt control, and 48.2% (P = 0.0012, n = 3) and 44.1% (P = 0.0036, n = 3) reduction in GE when compared with non-induced or induced R-Clover, respectively. Induction of R-Clover expression did not significantly affect measured GE (P = 0.7931, n = 3) when compared with non-induced control. (B) IFU assays conducted with samples normalized to GE showed a 49.6% (P < 0.0001, n = 3) and 13.6% (P = 0.0428, n = 3) reduction in recovered IFUs for induced R-dksACt and R-Clover, respectively, when compared with non-induced controls (set to 100%). The number of recovered IFUs following induction of R-dksACt was 36% less than that observed with induced R-Clover (P = 0.0004, n = 3). P-values were calculated using one-way analysis of variance (ANOVA) with Tukey's post hoc test.

DksA Ct retains major structural characteristics of DksA proteins
Amino acid sequence identity is commonly used as an indicator for predicting conservation of protein function (Tian and Skolnick 2003, Capra and Singh 2007, Krissinel 2007. Amino acid sequence alignment of DksA from various chlamydial species and multiple orthologs for which the crystal structures have been determined (DksA Ec , DksA Pa and DksA Af ) indicated high sequence conservation within the Chlamydia genus, and low sequence identity between the Chlamydia proteins and those encoded by genes in other bacteria (Fig. 3). Among humanadapted Chlamydia species, dksA Ct showed the highest identity with C. pneumoniae (90% identity at the amino acid level). In addition to apparent high conservation across the Chlamydia genus, homologs of dksA can also be found in nonpathogenic Protochlamydia amoebophila UWE25 and Protochlamydia naegleriophila KNic (Clark et al. 2016). Despite low (∼10%) primary sequence identity (Perederina et al. 2004, Rutherford et al. 2007, the transcription elongation factor GreA (GreA Ec ) has been shown to assume the physiological function of DksA in E. coli dksA ppGpp 0 mutants (Vinella et al. 2012), indicating that sequence identity does not consistently correlate with conserved function in this family of transcription factors.
Sequence analyses of DksA proteins from several bacteria has shown that while extensive diversity exists in the primary amino acid sequences, all proteins contain a conserved four-cysteine (C4) zinc-finger motif, CxxC-(x17)-CxxC, or a two-cysteine (C2) or one-cysteine (C1) zinc-independent motif, CxxS/T-(x17)-C/S/TxxA, with the first cysteine always conserved (Perederina et al. 2004, Henard et al. 2014. While the C4 proteins are sometimes referred to as DksA and the C2 and C1 proteins as DksA2 (Furman et al. 2013), here we will refer to all three groups as DksA proteins. The genomes of Chlamydia species all contain a single less frequently observed C1-DksA ortholog. Structures have been determined for two C4-DksA proteins, Escherichia coli (DksA Ec ) (Perederina et al. 2004) and Agrobacterium fabrum (DksA Af ), and one C2-DksA protein, P. aeruginosa (DksA Pa ) (Furman et al. 2013). Prototypical DksA proteins contain two domains that interact with RNAP: an N-terminal, two helix, coiled-coil domain that has been shown to interact with the secondary channel of RNAP (Molodtsov et al. 2018) and a C-terminal globular domain containing the Cx-motif plus a C-terminal αhelix that jointly interact with the β rim helices outside the RNAP secondary channel (Perederina et al. 2004, Lennon et al. 2012, Parshin et al. 2015. This interaction has high specificity, and it has been demonstrated in E. coli that a single point mutation in the C-terminal α-helix can result in loss of both binding and activity in vitro (Parshin et al. 2015). To facilitate a better general understanding of C1-DksA proteins and further elucidate the significance of widespread DksA conservation in Chlamydia (Domman and Horn 2015), we solved the 3D crystal structure of DksA Ct .
Despite the relatively low resolution (2.95Å) of the X-ray diffraction data, the final model had excellent refinement statistics (Table 1). Two protein molecules are observed in the asymmetric unit with the structure of each protomer having a backbone atom (N-C α -C = O) root-mean-standard deviation of 0.33Å between both chains (Fig. 4A). Superposition of the protein structures show that the topology of all four DksA protein structures are similar with four α-helices organized into two domains. The relative orientation of the two domains overlap in all four structures despite DksA Pa and DksA Ct containing two or one cysteine, respectively, and no zinc ion, further establishing that zinc is not required for function of DksA proteins (Furman et al. 2013). Instead, the cysteine and zinc content of DksA may impact its capacity to respond to reactive oxygen and nitrogen species (Henard et al. 2014, Crawford et al. 2016. Because all pathogenic chlamydiae appear to express a highly homologous C1-form of DksA, the protein may have evolved in these pathogens to maintain a functional structure in (intracellular) environments where zinc is not abundant. What is different between the four structures is the region towards the turn between α1 and α2 in the coiled coil domain. In the crystal structure of DksA Ct (C1-DksA), electron density is missing (E27-V62) and in the Nuclear Magnetic Resonance (NMR) structures for DksA Af (C4-DksA) there is no convergence of the calculated structures (K29-L58). These observations in crystal (Oldfield et al. 2013) and NMR (Konrat 2014) structures may be correlated to intrinsically disordered regions. On the other hand, the loop between α1 and α2 is significantly smaller for DksA Pa (chain C, R50-A58) and DksA Ec (F69-P70), averaging eight and two residues, respectively. To explore the possibility that this difference could be attributed to varying degrees of intrinsic disorder we analyzed all four primary amino acid sequences of the proteins depicted in Fig. 4A using IUPred2A (Mészáros et al. 2018, Erdős andDosztányi 2020). Alignment of these sequences starting with the first residue of the first N-terminal α-helix identifies a region of ∼20 residues between the first two N-terminal α-helices with IUPred scores above 0.5, the threshold for disorder, for three of the four proteins (∼10 residues for DksA Pa showed scores below the cutoff) (Fig. S3, Supporting Information). We speculate that in monomeric DksA proteins the tip of the coiled coil domain is disordered with the helical regions at the C-terminal and N-terminal ends of α1 and α2, respectively, being transient in nature. This structural fluidity may be necessary for this domain to dock with the secondary channel of RNAP  (Garner et al. 1999). These regions are more helical in the crystal structures of DksA Pa and DksA Ec perhaps only due to crystal packing contacts that restrict motion and stabilize the helical state (Oldfield et al. 2013). Figure 4B is a cartoon representation of one of the two structures in the asymmetric unit showing that the protein contains four α-helices with the N-terminal two helices forming a coiled coil domain and the two C-terminal helices part of a C-terminal globular domain containing a C1-motif. In both chains, electron density was insufficient to model the eight residues of the Nterminal tag, one or two residues at the C-terminus, and the region between L26(A)/E27(B) and V62, residues between the two helices of the coiled coil domain. No electron density corresponding to zinc or any other metal was observed.

DksA Ct does not functionally complement DksA Ec
Complementation of dksA-deficient E. coli has been conducted successfully with genes encoding C4-, C2-and C1-DksA proteins (Blaby-Haas et al. 2011, Pal et al. 2012, Lennon et al. 2014. For example, the C1-DksA ortholog of the α-proteobacterium R. sphaeroides shares 42% identity with DksA Ec , and has been shown to function alone and synergistically with ppGpp (Lennon et al. 2014). Divergent primary sequences, unresolved structural domains and the unique biology of C. trachomatis prompted analysis to determine whether dksA Ct can functionally complement the prototypical E. coli ortholog dksA Ec. DksA functionality can be tested based on the role of DksA in regulating amino acid synthesis in E. coli during growth on minimal M9 medium (Jude et al. 2003). dksA complementation in E. coli has shown that the dksA genes of Vibrio cholera (Pal et al. 2012), R. sphaeroides (Lennon et al. 2014) and P. aeruginosa (Blaby-Haas et al. 2011) can functionally complement dksA Ec . In our hands, P. aeruginosa dksA (dksA1 Pa ) resulted in partial rescue of an E. coli mutant unable to express dksA Ec , while complementation with dksA Ct did not restore the growth defect (Fig. 5). Moreover, complementation with P. aeruginosa dksA2 (dksA2 Pa ) did not result in recovery of the growth defect exhibited by E. coli lacking the dksA gene. Growth in the absence of inducer is likely a result of leaky expression of DksA Ec in the dksA deletion mutant. Overall, dksA Ct does not have activity complementary to that of dksA Ec or dksA1 Pa suggesting an alternative binding motif or function for dksA Ct compared with that of prototypical DksA proteins.

dksA Ct expression is regulated from promoter sites both upstream of and within nrdR
Gene linkage and co-expression can be related to the function of two or more genes in a coordinated biological process. In all chlamydial genomes analyzed, dksA is located immediately downstream of nrdR, encoding the NrdA/B repressor NrdR, a negative regulator of deoxyribonucleotide synthesis. lspA, encoding a lipoprotein signal peptidase, is located immediately downstream of dksA Ct . The genomic localization and organization of dksA with nrdR and lspA in chlamydial genomes, and intergenic regions of only 5-24 bp, are consistent with their control by a single promoter upstream of nrdR. This conserved orientation provides a potential link between the role of dksA and the metabolic function of nrdR in nucleic acid metabolism. Previously published deep sequencing data indicate that nrdR, dksA and lspA in C. pneumoniae are transcribed as a polycistronic mRNA (Albrecht et al. 2011) and suggest that the same may be true in C. trachomatis (Albrecht et al. 2010).
To determine whether dksA Ct could be regulated from an independent promoter downstream of the promoter controlling expression of the nrdR-dksA Ct -lspA operon, C. trachomatis was transformed with transcriptional reporter constructs based on the pBOMB vector (Bauler and Hackstadt 2014) containing sequences 830 or 469 bp upstream of dksA Ct linked to GFP-LVA (Fig. 6A). Constitutive expression of mCherry was used as a positive control for transformation. GFP was detected regardless of construct configuration suggesting that dksA Ct can be transcribed from promoters both upstream of and within nrdR (Fig. 6B). No signal was detected from the empty vector control.
To independently verify the existence of two unique transcripts containing dksA Ct , total RNA was extracted from C. trachomatis-infected HeLa cells at 20 and 30 hpi and northern blotting performed with oligonucleotide probes complementary to dksA Ct . Blots indicate that dksA Ct is expressed as two transcripts with a prominent band at ∼1.0 kb and a less prominent band at 1.5 kb (Fig. 7A). This pattern is consistent with polycistronic regulation via the 1.5 kb fragment, as the distance from the published TSS of nrdR to the lspA stop codon is 1477 bp (Albrecht et al. 2010). Characterization of the 1.0 kb fragment required additional verification due to the similar sizes of nrdR  (465 bp) and lspA (504 bp). Therefore, we utilized neural networkbased prediction software (Reese 2001) to search for sites within the identified operon region that may constitute an alternative promoter. Four sites were identified, one of which correlated directly with the TSS previously determined for nrdR. Of the other three predicted TSSs, two were located within nrdR and one within dksA Ct . Using the predictions as a guide, 5 RACE was conducted using gene specific primers (GSPs) designed to bind within dksA Ct (Fig. 7B; Fig. S4, Supporting Information). Sequencing of 5 RACE fragments indicate a promoter site ∼80-150 bp upstream of the dksA Ct ORF, which directly correlates with the 783880-783925 region predicted to contain the alternative promoter.
Transcript levels for dksA Ct and the adjacent genes nrdR and lspA were determined by RNA-Seq at time points correlating with early, mid and late stages of the chlamydial developmental cycle (Fig. 7C). Data obtained by RNA-Seq support independent regulation of dksA Ct through a 5-6-fold increase in dksA Ct reads when compared with nrdR. Additionally, the expression pattern of DksA Ct (Fig. 1) correlates with the temporal pattern of transcript abundance observed in the RNA-Seq data set. As a final confirmation, quantitative reverse transcription polymerase chain reaction (RT-qPCR)-based transcriptional analysis conducted using cDNA extracted at 30 hpi indicated a 3-4-fold increase in dksA Ct transcript when compared with a region of nrdR upstream from the 5 RACE site (Fig. S5, Supporting Information). This increase in transcript abundance corroborates data obtained by RNA-Seq, further supporting transcriptional regulation for dksA Ct from two independent promoters.

Conclusions
DksA has been described as one of eight transcription factors conserved across the phylum Chlamydiae (Domman and Horn Figure 7. dksACt is expressed as two distinct transcripts. Northern blot analysis was used to confirm expression of dksACt as two distinct RNA species. (A) Denaturing gel stained with EtBr and corresponding membrane hybridized with a biotinylated probe complementary to a segment of dksACt visualized using chemiluminescence. Total RNA extracted from noninfected HeLa cells was used as a control for nonspecific binding of the probe to host RNA. Lane 1: RNA from uninfected HeLa cells. Lane 2: RNA isolated from C. trachomatis-infected HeLa cells, 20 hpi (n = 2). Lane 3: RNA isolated from C. trachomatis-infected HeLa cells, 30 hpi (n = 3). All lanes were loaded with 15 μg RNA. (B) Schematic showing the relative locations for dksACt-specific primers (a, b and c) used to determine the dksACt promoter start site within nrdR. The number of detected matching sequences and genomic location within the C. trachomatis genome are indicated. The target site for the biotinylated oligonucleotide probe within dksACt is also shown. (C) A previously generated RNA-Seq data set (Grieshaber et al. 2018) was used to assess whether genes in the nrdR-dksA-lspA operon show temporal differences in expression (n = 4). 2015). Based upon data from previously studied C4-, C2-and C1-DksA orthologs, we predicted that DksA Ct is maintained as a protein associated with RB-EB transitions in the chlamydial developmental cycle. Herein, we showed that DksA Ct is expressed maximally at the RB-EB transition point (i.e. 15-20 hpi), a unique expression pattern compared with the stable expression observed for DksA Ec (Paul et al. 2004, Brown et al. 2002. Ectopic overexpression of DksA Ct prior to RB-EB transition produced a defect in both replication and EB generation. We found that dksA Ct does not functionally complement dksA Ec with respect to recovery of replication during growth on minimal medium. DksA Ct has an overall domain structure similar to other C2-and C4-type DksA proteins despite containing only one cysteine residue. Upon comparing our C1-type DksA structure to the structure of other C2-and C4-type DksA proteins we speculate that the 'tip' of the coiled coil domain may be transiently structured/intrinsically disordered to assist binding to the secondary channel of RNAP. Expression of DksA Ct is likely regulated by tandem promoter regions, a regulatory scheme also described for other genes in this pathogen (Lambden et al. 1990, Rosario andTan 2012). The phenotype resulting from induced DksA Ct expression prior to RB-EB transition and the correlation between dksA Ct and nrdR expression is consistent with a role for DksA Ct in the C. trachomatis developmental cycle. It should be mentioned that DksA can function in ways that could be beneficial to C. trachomatis even in the absence of a stringent response, including resolution of transcription conflicts (Tehranchi et al. 2010) and protection of phleomycin-induced double-stranded breaks in DNA (Sivaramakrishnan et al. 2017). Therefore, selective pressure to maintain dksA in chlamydiae could be related to such alternative functions.
It will be valuable to pursue loss of function phenotypes in future studies via deletion of dksA Ct (Mueller et al. 2017, Keb et al. 2018, Keb and Fields 2020 or CRISPR interference (Ouellette 2018, Ouellette et al. 2021 in case dksA Ct is essential to C. trachomatis intracellular replication and/or developmental transitions. Mechanistic understanding of how DksA Ct functions in C. trachomatis biology will require analysis of whether the protein directly influences pathogen gene expression by binding to RNAP akin to prototypical DksA proteins, or whether DksA Ct is associated with a different mechanism of action, such as interfering with the binding of another protein to RNAP, thus indirectly affecting transcriptional regulation.

Maintenance, purification and quantification of Chlamydia
Chlamydia trachomatis RB and EB cell forms were purified by density gradient centrifugation as previously described (Grieshaber et al. 2018). Chlamydial EBs were stored at −80 • C in sucrosephosphate-glutamate (Bovarnick et al. 1950) or K36 (Weiss 1965) buffer until use. Crude preparations of EBs for use in transformations were generated as previously described (Mueller et al. 2017). Chlamydia trachomatis was quantified by measurement of GE by SYBR green-based qPCR using a CFX96 or CFX384 real time PCR Detection System (BioRad, Hercules, CA) and primers targeting the hctA gene of C. trachomatis.

Plasmid construction and transformation of C. trachomatis
PCR primers used in this study are listed in Table S1 (Supporting Information). The vector pBOMB3cdMCI::incD-GFPLVA (Bauler and Hackstadt 2014) was linearized by PCR to remove the incD cassette using Q5 High-Fidelity DNA polymerase (New England Biolabs; Ipswich, MA). For dksA transcriptional fusion constructs, two inserts were generated spanning regions of 469 and 830 base pairs upstream from the annotated start site of dksA. For the E-Riboswitch dksA construct, dksA was inserted into pBOMB4 with an E-Riboswitch (Topp et al. 2010) driven by a T5 promoter to generate pBOMB4::E-Riboswitch-dksA Ct . All DNA inserts were PCRamplified using Phusion DNA polymerase (New England Biolabs; Ipswich, MA) and primer sets were generated according to the In-Fusion HD Cloning (Takara Bio, USA) guidelines. Linearized pBOMB4 and pBOMB4::GFPLVA were DpnI treated, then ligated to each insert and transformed into Stellar competent cells using the In-Fusion HD Cloning Kit (Takara Bio, USA). After incubation overnight at 37 • C on LB agar plates containing 100 μg/mL carbenicillin (LB Carb ) (Chem-Impex International; Wood Dale, IL), colonies were selected for replating and screening via PCR. Plasmids were isolated using the Purelink Quick Plasmid Miniprep Kit (Invitrogen; Carlsbad, CA) and sequenced for insert confirmation. The confirmed plasmids were transformed into methyltransferase-deficient E. coli K-12 ER2925 competent cells and incubated overnight. Positive colonies were subcultured in 50 mL LB Carb broth at 37 • C, and unmethylated plasmids were isolated using the GenElute HP Endotoxin Free Plasmid Maxiprep Kit (MilliporeSigma; St Louis, MI). Eluted plasmids were either concentrated to ∼500 ng/μL using the DNA Clean and Concentrator-100 kit (Zymo Research; Irvine, CA) or via sodium acetate and isopropanol-mediated DNA precipitation. Unmethylated plasmids purified from E. coli were transformed into C. trachomatis essentially as described (Mueller et al. 2017). For some transformations of C. trachomatis, density gradient purified EBs (1 × 10 8 GE/well) were mixed with 2 μg of plasmid DNA. Cultures identified to have positive transformants as determined by fluorescence microscopy, were passaged two additional times to remove residual penicillin G susceptible bacteria and increase the titer of transformed cells. Clonal populations were generated via extraction of C. trachomatis from inclusions by micromanipulation repeated four times under limiting dilution. Plasmid DNA was obtained from isolated clones and transformed into E. coli where five colonies expressing mCherry were sequenced for confirmation of clonality.

Sequence analysis
Amino acid sequence alignments were done using the ClustalW (Thompson et al. 1994) alignment tool using MEGA software (Tamura et al. 2021) and the MView multiple alignment viewer (Brown et al. 1998).

Analysis of DksA Ct expression
Protein lysates for analysis of DksA Ct expression by western blotting were obtained from 10-15 HeLa cell cultures (T-175) infected with C. trachomatis at an multiplicity of infection (MOI) of 300; bacteria were enriched by density gradient centrifugation using a 30% diatrizoate meglumine (MD-76R) pad essentially as described (Howard et al. 1974, Grieshaber et al. 2018. Bacteria were enumerated by analysis of GE, then lysed in Laemmli buffer containing β-mercaptoethanol (Bio-Rad; Hercules, CA) and heat denatured at 100 • C for 10 min. Twelve % sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gels were loaded with the equivalent of 2 × 10 8 GE of each sample per well and electrophoresed at 25 mA for 40-60 min, then transferred to a 0.45 um PVDF membrane using a Trans-Blot Turbo transfer apparatus (Bio-Rad; Hercules, CA). The membrane was blocked in PBS + 0.1% Tween-20 (PBS-T) and 3% nonfat dry milk overnight at 4 • C. The membrane was probed with a custom protein-A purified anti-DksA Ct antibody raised in rabbits against a truncated recombinantly expressed DksA Ct antigen in combination with a goat anti-rabbit poly-HRP secondary antibody (Invitrogen; Carlsbad, CA). The membrane was treated with the SuperSignal West Femto luminol and peroxide solution (ThermoFisher; Waltham, MA) and imaged using a ChemiDoc Imaging System (Bio-Rad; Hercules, CA). Animal use was approved by the Institutional Animal Care and Use Committee at Washington State University.

RNA extraction and northern blotting
HeLa cells were established in T-175 cell culture flasks and infected at an MOI of 300. At 20 and 30 hpi, cultures were disrupted by scraping in 1 mL K-36 buffer (Bovarnick et al. 1950) and centrifuged at 100 × g at 4 • C for 3 min to remove large particles. Bacteria were pelleted from the supernatant via centrifugation at 20 000 × g at 4 • C for 30 min. Pellets were then resuspended in 1 mL TRIzol reagent (Invitrogen, USA). HeLa RNA (control) was collected by scraping uninfected confluent monolayers in 4 mL TRIzol reagent. Samples were homogenized using a FastPrep 24 Tissue Homogenizer (MP Biomedicals; Santa Ana, CA) with 0.1 mm zirconia beads at 4 m/s for 20 s, repeated two times. Total RNA was extracted using chloroform, isopropanol and ethanol according to the manufacturer's protocol. RNA quantity was assessed using a Nanodrop Spectrophotometer (ThermoFisher; Waltham, MA), and RNA integrity measured using an Agilent 2100 Bioanalyzer (Agilent Technologies; Santa Clara, CA) and the RNA 6000 Nano Kit according to manufacturer's instructions.
Northern blotting of isolated total RNA was done using the NorthernMax Kit (Invitrogen, USA). PCR-generated dksA dsDNA was used as a positive control for detection. A custom 5 Biotin tagged ssDNA oligonucleotide probe (Table S1, Supporting Information) was designed to bind dksA mRNA according to the ULTRAhyb Oligo Buffer (Invitrogen; Carlsbad, CA) recommendations. The RiboRuler high range RNA ladder and 2× loading dye (ThermoFisher; Waltham, MA) were used to assess mRNA size. The membrane was cut to match the exact size of the denaturing gel, and RNA transferred via capillary action for 2 h. The membrane was incubated in a UV crosslinker (Hoefer; Holliston, MA) for 1 min, repeated 3 times. The membrane was pre-hybridized with rotation in a 50 mL conical tube for 1 h in ULTRAhyb Oligo Buffer at 40 • C. The probe was added to 3.5 mL buffer at a concentration of ∼600 pM and hybridized at 40 • C for 17-20 h. The membrane was washed in the NorthernMax low stringency solution twice for 5 min at RT, and once at hybridization temperature for 2 min. The washed membrane was treated with the Chemiluminescence Nucleic Acid Detection Module Kit (ThermoFisher; Waltham, MA) according to the manufacturer's instructions, and imaged using a ChemiDoc Imaging System (Bio-Rad; Hercules, CA).

RACE
RNA isolated at 24 hpi was converted into cDNA tagged with the SMARTer II A Oligonucleotide using random hexamers as described in the SMARTer RACE 5 /3 Kit (Takara Bio USA; Mountain View, CA). cDNA libraries were diluted as needed, and 5 RACE PCR reactions were run using three separate GSPs designed to target the intragenic dksA Ct sequence (Table S1, Supporting Information). PCR products were electrophoresed on a 1% TAE gel stained with SYBR Safe DNA Gel Stain (Invitrogen; Carlsbad, CA), and bands were excised for DNA isolation using the Nucleospin Gel and PCR Cleanup Kit (Macherey-Nagel; Düren, Germany). Fragments were then cloned using the Clone-JET PCR cloning kit (ThermoFisher; Waltham, MA) and plasmids were isolated from positive transformants using the Nucleospin Plasmid Kit (Macherey-Nagel; Düren, Germany). Plasmids were sequencing using pJET1.2 vector specific primers (Eurofins Genomics). Clones were analyzed using NCBI BLAST against the C. trachomatis serovar L2 genome. Sequences not containing both the GSP and Universal Primer Mix regions, indicative of incomplete sequences, were excluded from the analysis.

PCR and RNA-Seq
cDNA for both conventional PCR analysis of polycistronic RNA and RT-qPCR applications was generated using the SuperScript III First-Strand Synthesis kit (Invitrogen; Carlsbad, CA) with RNA isolated at 20 and 30 hpi. Negative controls not containing reverse transcriptase were generated for each time point. Primers specific to nrdR and lspA (Table S1, Supporting Information) were utilized in conventional PCR to amplify cDNA for operon determination. For transcriptional analysis, C. trachomatis serovar L2 genomic DNA was extracted from density gradient purified EBs using phenol/chloroform precipitation. A dilution series of genomic DNA was generated and used for calculation of primer efficiencies and for calculation of relative copy number. Primer efficiencies between 85% and 110% were deemed acceptable for direct comparison of transcript levels. RNA-Seq data was recovered from a previously published data set (Grieshaber et al. 2018).

Ectopic expression of DksA in C. trachomatis
HeLa cells were grown to confluency in T-25 cell culture flasks. The culture medium was then replaced with RPMI-1640 supplemented with 10% (v/v) FetalPlex and 10 μg/mL Gentamicin. Each condition was tested in duplicate with two cultures supplemented with 0.5 mM theophylline and the other two serving as controls. Theophylline was added to the culture medium prior to inoculation. To facilitate comparative analysis, each flask was inoculated to reach equivalent levels (50-75%) of infection using bacteria carrying the pBOMB4::E-Riboswitch-dksA Ct or p2TK2-SW2::E-Riboswitch-Clover (Grieshaber et al, 2022) vectors and incubated at 37 • C in a 5% CO 2 incubator for 30 h. Induction of pBOMB4::E-Riboswitch-dksA Ct results in expression of an untagged DksA Ct protein. The medium was then removed, and HeLa cell monolayers were washed with K-36 buffer. Infected host cells were disrupted by vigorous scraping in K-36 buffer and then host cell debris was removed from the mixture via centrifugation at 100 × g for 3 min at 4 • C. The supernatant was removed, and Chlamydia was pelleted at 20 000 × g for 15 min at 4 • C. Pelleted bacteria were resuspended in 30 μL SPG and frozen at −80 • C. GEs were enumerated using qPCR and IFU assays were conducted using HeLa cells grown to confluency in 12-well plates with inocula normalized to GE.

Microscopy
Micrographs were acquired using a Leica DMi8 inverted microscope (Leica Microsystems; Buffalo Grove, IL) equipped with an X-Cite LED light source and DMC2900 camera. Chlamydia trachomatis IFU assays were done using methanol fixed samples stained with a fluorescein isothiocyanate-tagged anti-MOMP antibody (PA1-73073 Thermo Fisher; Waltham, MA) at 1/250 dilution in PBS buffer. Images were analyzed and processed using ImageJ (The National Institute of Health; Bethesda, MD). Adjustments of contrast or signal intensity were applied to the entire image.

Cloning of dksA Ct
The dksA Ct gene (Gene ID: 5858340) was amplified from the genomic DNA of C. trachomatis serovar L2 (LGV 434/Bu) (NCBI: 471472) and inserted into the expression vector BG1861 (a derivative of pET14b) (Myler et al. 2009) at a site containing an uncleavable, 8-residue N-terminal tag (MAHHHHHH-). Using a heat shock method, the recombinant plasmid was used to transform E. coli BL21(DE3)-R3-pRARE2 cells (gift from SGC Toronto). From these transformed cells, stocks were prepared (∼1 mL LB media, OD 600 ∼0.8) from a single colony and stored at −80 • C in glycerol (∼15%) solution until preparation of samples for structural analysis.
Crystallization, X-ray data collection and structure refinement of DksA Ct For X-ray crystallographic analysis, DksA Ct was expressed and purified following standard SSGCID protocols (Bryan et al. 2011). The protein was concentrated to 24.6 mg/mL in 25 mM HEPES, 500 mM NaCl, 5% (v/v) glycerol, 2 mM DTT, and 0.025% sodium azide (v/v), pH 7.5. Initial crystallization conditions were searched for using 0.4 (protein) plus 0.4 (precipitant) μL drops in XJR crystallization trays (Rigaku Reagents; Bainbridge Island, WA) and several commercial screens: JCSG+, JCSG-Top96, Wizard 1/2, Wizard 3/4 (Rigaku Reagents; Bainbridge Island, WA), Crystal Screen, IndexHR (Hampton Research; Aliso Viego, CA), MCSG-1, MCSG-2 (Microlytic/Anatrace; Maumee, OH), Morpheus 1 and 2, PACT (Molecular Dimensions; Holland, OH). Initial crystal hits were optimized starting from CrystalScreen, condition B5: 200 mM lithium sulfate, 25% (w/v) PEG 4000, 100 mM Tris, pH 8.5. The crystals were cryoprotected with 15% ethylene glycol and vitrified in nylon loops by plunging into liquid nitrogen. Diffraction data up to 2.95Å resolution were collected at the Advanced Photon Source (APS), Live Sciences Collaborate Access Team (LS-CAT) beamline 21-ID-F, at a wavelength of 0.97872Å with a Rayonix MX-300 detector. Diffraction data statistics are summarized in Table 1. The structure could not be solved using the standard SSGCID molecular replacement strategies. However, with MR-Rosetta (Terwilliger et al. 2012) as implemented in Phenix (Adams et al. 2011), a dimer solution could be found that was based on PDB entries 4IJJ A, 1TJL H, 2KQ9 A, 2KGO A and 5W1S N. The structure was then completed with iterative refinement cycles in phenix.refine (Zwart et al. 2008), and real space modeling in Coot (Emsley et al. 2010). The quality of the model was assessed with tools built into Coot and phenix.refine, such as MolProbity (Williams et al. 2018). The coordinates and structure factors were deposited in the PDB with code 6PTG. Diffraction images are available at proteindiffraction.org.

Statistical analysis
Statistical analyses were conducted using Prism (GraphPad Inc.; San Diego, CA) software. Statistical analysis and normalization of read counts for RNA-Seq data (Fig. 7) was done using DESeq2 in R (Love et al. 2014). Normalization from Dseq2 was used for transcript visualization in Integrated Genomics Viewer (Robinson et al. 2011).