Role for GrgA in Regulation of σ²⁸-Dependent Transcription in the Obligate Intracellular Bacterial Pathogen Chlamydia trachomatis

Chlamydia trachomatis is the number one sexually transmitted bacterial pathogen worldwide. A substantial proportion of C. trachomatis-infected women develop infertility, pelvic inflammatory syndrome, and other serious complications. C. trachomatis is also a leading infectious cause of blindness in underdeveloped countries. The pathogen has a unique developmental cycle that is transcriptionally regulated. The discovery of an expanded role for the Chlamydia-specific transcription factor GrgA helps us understand the progression of the chlamydial developmental cycle.

ABSTRACT

The obligate intracellular bacterial pathogen Chlamydia trachomatis has a unique developmental cycle consisting of two contrasting cellular forms. Whereas the primary Chlamydia sigma factor, σ⁶⁶, is involved in the expression of the majority of chlamydial genes throughout the developmental cycle, expression of several late genes requires the alternative sigma factor, σ²⁸. In prior work, we identified GrgA as a Chlamydia-specific transcription factor that activates σ⁶⁶-dependent transcription by binding DNA and interacting with a nonconserved region (NCR) of σ⁶⁶. Here, we extend these findings by showing GrgA can also activate σ²⁸-dependent transcription through direct interaction with σ²⁸. We measure the binding affinity of GrgA for both σ⁶⁶ and σ²⁸, and we identify regions of GrgA important for σ²⁸-dependent transcription. Similar to results obtained with σ⁶⁶, we find that GrgA's interaction with σ²⁸ involves an NCR located upstream of conserved region 2 of σ²⁸. Our findings suggest that GrgA is an important regulator of both σ⁶⁶- and σ²⁸-dependent transcription in C. trachomatis and further highlight NCRs of bacterial RNA polymerase as targets for regulatory factors unique to particular organisms.

IMPORTANCE Chlamydia trachomatis is the number one sexually transmitted bacterial pathogen worldwide. A substantial proportion of C. trachomatis-infected women develop infertility, pelvic inflammatory syndrome, and other serious complications. C. trachomatis is also a leading infectious cause of blindness in underdeveloped countries. The pathogen has a unique developmental cycle that is transcriptionally regulated. The discovery of an expanded role for the Chlamydia-specific transcription factor GrgA helps us understand the progression of the chlamydial developmental cycle.

INTRODUCTION

Each year, ∼60% of the 2.2 million cases of notifiable infections reported to the Centers for Disease Control and Prevention (CDC) are due to the sexually transmitted pathogen Chlamydia trachomatis (1, 2). In addition, because the infection is mostly asymptomatic, the CDC estimates only 1 of 10 people infected with C. trachomatis is reported (3). Nonetheless, without antibiotic treatment, the infection often leads to serious complications, such as infertility and pelvic inflammatory syndrome in women. Furthermore, some C. trachomatis serotypes cause ocular infection and are still the most common infectious microbes associated with blindness in underdeveloped countries (4, 5).

Like other chlamydiae, C. trachomatis is an obligate intracellular Gram-negative bacterium that exists in two cellular forms with contrasting properties (6). The small elementary body (EB) is infectious and capable of extracellular survival but is incapable of proliferation. Following binding to a cellular receptor(s), the EB enters a host cell membrane-derived vacuole through endocytosis (7). Within the vacuole, termed inclusion, the EB differentiates into a larger cellular form, termed reticulate body (RB), within several hours. No longer infectious, the RB divides exponentially by binary fission until around 20 h, when a significant portion of RBs redifferentiate back into EBs, while some RBs continue to proliferate (8). Progeny EBs along with residual RBs are released from infected cells following cell lysis. Alternatively, whole inclusions may be released from infected cells (9).

The 1-million-bp C. trachomatis genome carries fewer than 1,000 genes (10). Microarray analyses demonstrated that the majority of these genes are transcribed starting a few hours postinoculation throughout the remaining developmental cycle, whereas a small number of genes are transcribed immediately following cell entry and another small set of genes are transcribed only at late stages (11, 12). Transcriptome sequencing (RNA-seq) detected distinct sets of gene transcripts specifically enriched in either EBs or RBs (13), and purified EBs and RBs have been found to transcribe different sets of genes in axenic media (14). These findings suggest that the progression of the chlamydial developmental cycle is transcriptionally regulated.

Transcription is initiated following binding of the RNA polymerase (RNAP) to the gene promoter (15). The bacterial RNAP holoenzyme is comprised of the catalytic core enzyme and a σ factor, which is required for promoter recognition (16). Transcription of the vast majority of C. trachomatis genes involves σ⁶⁶, a homolog of σ⁷⁰ that is often referred to as the housekeeping σ factor in eubacteria (16). Expression of some (but not all) chlamydial late genes depends on σ²⁸. Several genes possess both a σ⁶⁶ promoter and a σ²⁸ promoter (17).

GrgA (with gene codes CT_504 and CTL0766 for C. trachomatis serovars D and L2, respectively) is a Chlamydia-specific transcription activator (18). It was identified as a protein bound to the σ⁶⁶-dependent promoter of defA, which encodes peptide deformylase, an enzyme required for bacterial protein maturation and regulated protein degradation. In addition to defA, a midcycle gene, GrgA also stimulates transcription from another midcycle promoter (ompA), an early promoter (rRNA P1), and a late promoter (hctA), suggesting that GrgA functions as a general activator of σ⁶⁶-dependent genes (18). In this report, we demonstrate that GrgA also stimulates σ²⁸-dependent gene transcription in vitro. Thus, our findings suggest that GrgA plays an expanded role in gene expression during the C. trachomatis developmental cycle as a regulator of both σ⁶⁶- and σ²⁸-dependent transcription.

RESULTS

GrgA physically interacts with σ²⁸.

To assess whether GrgA potentially regulates expression of σ²⁸-dependent genes, we determined whether GrgA can interact with σ²⁸. We performed protein pulldown assays using differential epitope tagging. The Strep-Tactin beads, which have affinity for the Strep-tag (19), precipitated NH-σ²⁸ (N-terminally poly-His-tagged C. trachomatis σ²⁸) in a manner that was dependent on the N-terminally Strep-tagged GrgA (NS-GrgA) (Fig. 1A). Reciprocally, NH-GrgA was pulled down in an NS-σ²⁸-dependent manner (Fig. 1B). The results establish that GrgA can directly interact with σ²⁸.

FIG 1.

GrgA physically interacts with σ²⁸. (A) Pulldown of NH-σ²⁸ by Strep-Tactin bead-immobilized NS-GrgA. Shown is a Western blot detecting NH-σ²⁸. (B) Pulldown of NH-GrgA by Strep-Tactin bead-immobilized NS-σ²⁸. Shown is a Western blot detecting NH-GrgA.

GrgA has a lower affinity for σ²⁸ than for σ⁶⁶.

We next determined the binding affinities of GrgA for both σ²⁸ and σ⁶⁶. We first compared the efficiencies of the σ factors in GrgA binding by performing competitive pulldown assays. As expected, NS-GrgA efficiently pulled down NH-σ²⁸ and CH-σ⁶⁶ in separate reactions (see Fig. S1 in the supplemental material). However, in the presence of equal molar concentrations of NH-σ²⁸ and CH-σ⁶⁶, NS-GrgA pulled down only CH-σ⁶⁶ but not NH-σ²⁸ (Fig. S1), indicating that GrgA has a lower affinity for σ²⁸ than σ⁶⁶.

We quantitatively characterized GrgA binding by σ²⁸ and σ⁶⁶ with biolayer interferometry (BLI) using the BLItz system, which detects light wavelength shifts at the biosensor tip with an immobilized ligand following binding of an analyte in a real-time manner (20). Whereas representative BLItz recordings using NH-GrgA as a ligand and either CS-σ⁶⁶ or NS-σ²⁸ as an analyte are shown in Fig. S2A and B, values of kinetic parameters are provided in Table 1. The CS-σ⁶⁶ analyte yielded a statistically highly significant 25-fold higher association rate constant (k_a) than that of the NS-σ²⁸ analyte, suggesting that CS-σ⁶⁶ binds NH-GrgA much faster than NS-σ²⁸. CS-σ⁶⁶ also demonstrated a 3-fold statistically significant increase in disassociation rate constant (k_d), suggestive of moderately higher dissociation from NH-GrgA. Compared to the NH-GrgA–CS-σ⁶⁶ interaction, the NH-GrgA–NS-σ²⁸ interaction had a 32-fold higher disassociation equilibrium constant (K_D), indicating that GrgA has a lower overall affinity for σ²⁸ than for σ⁶⁶.

TABLE 1.

GrgA binds σ⁶⁶ with a higher affinity than σ^28a

Ligand	Analyte	n	ka		kd		KD
			1/Ms	% control	1/s	% control	M	% control
NH-GrgA	CS-σ66	6	(1.9 ± 1.3) × 106	100	(9.3 ± 5.2) × 10−3	100	(6.9 ± 5.9) × 10−9	100
NH-GrgA	NS-σ28	8	(1.5 ± 1.7) × 104; P = 0.001	4.0	(2.8 ± 0.8) × 10−3; P = 0.001	30	(2.2 ± 1.1) × 10−7; P < 0.001	3,188
CH-σ66	NS-GrgA	4	(7.7 ± 1.3) × 105	100	(8.9 ± 0.6) × 10−3	100	(1.2 ± 0.1) × 10−8	100
NH-σ28	NS-GrgA	6	(2.1 ± 0.9) × 105; P < 0.001	27	(5.6 ± 1.2) × 10−2; P < 0.001	629	(3.4 ± 2.0) × 10−7; P = 0.013	2,833

^aBLItz assays were performed with His biosensors using the indicated ligand and analyte pairs. Representative graphs of recordings are shown in Fig. S2. Values of kinetic parameters (averages ± SDs) were generated by the BLItz Pro software (20). k_a (association rate constant) is defined as the number of complexes formed per second in a 1 M solution of ligand and analyte (1/Ms). k_d (disassociation rate constant) is defined as the number of complexes that decay per second (1/s). K_D (disassociation equilibrium constant), defined as the concentration at which 50% of ligand binding sites are occupied by the analytes, is k_d divided by k_a. n, number of experimental repeats. P values were calculated using 2-tailed Student's t tests.

We performed reciprocal BLI using CH-σ⁶⁶ and NH-σ²⁸ as ligands and NS-GrgA as the analyte to validate the difference in GrgA binding by the σ factors presented above (Table 1 and Fig. S2C and D). Consistent with the trend in k_a value changes presented above, the NS-GrgA analyte also demonstrated a statistically significantly higher k_a for CH-σ⁶⁶ than for NH-σ²⁸, although the difference is smaller (25-fold versus 3.7-fold). Interestingly, the k_d values reveal that NS-GrgA also dissociates from CH-σ⁶⁶ 6-fold slower than that from NH-σ²⁸. Compared to the CH-σ⁶⁶–NS-GrgA interaction, the NH-σ²⁸–NS-GrgA interaction had a 28-fold higher K_D, which is nearly identical to the 32-fold higher K_D detected for the NH-GrgA–NS-σ²⁸ interaction versus the NH-GrgA–CS-σ⁶⁶ interaction. Thus, competitive pulldown assays and BLI establish that GrgA has a lower affinity for σ²⁸ than for σ⁶⁶.

GrgA stimulates σ²⁸-dependent transcription.

To determine whether GrgA can stimulate σ²⁸-dependent transcription, we performed in vitro transcription assays using pMT1212, a transcription reporter plasmid carrying the promoter of a gene encoding a histone-like protein (hctB) in C. trachomatis (21). Consistent with previous findings (21), transcription from the hctB promoter required the addition of NH-σ²⁸ to the C. trachomatis RNAP (cRNAP) (Fig. 2A). Interestingly, GrgA demonstrated a dose-dependent stimulatory effect on the transcription from the promoter (Fig. 2B and C). These data suggest that GrgA can increase the expression of genes with a σ²⁸-dependent promoter in addition to genes with a σ⁶⁶-dependent promoter (18).

FIG 2.

GrgA stimulates σ²⁸-dependent transcription using C. trachomatis RNAP. (A) Transcription from the C. trachomatis hctB promoter in the pMT1212 report plasmid is dependent on the addition of NH-σ²⁸ to the chlamydial RNAP. (B) Gel image showing dose-dependent stimulation of transcription from the hctB promoter by GrgA. (C) Averages and SDs for three independent measurements are shown. (B and C) Molar ratios of GrgA to σ²⁸ were 0, 10, 20, and 33.

Residues 138 to 165 in GrgA are required for binding both σ²⁸ and DNA and for activating σ²⁸-dependent transcription.

A series of His-tagged GrgA deletion mutants (Fig. S3A and B) were tested for the effects on transcription from the hctB promoter (Fig. 3A). Noticeably, GrgAΔ114-165 was completely defective in activating transcription from the σ²⁸-dependent promoter, whereas GrgAΔ1-64 also demonstrated a significant 50% loss of transcription activation activity (Fig. 3A). Deletion of other regions (residues 65 to 113, 166 to 266, and 207 to 288) from GrgA had either no or minimal effects on transcription activation (Fig. 3A).

FIG 3.

Residues 138 to 165 in GrgA are required for binding both σ²⁸ and DNA and for activating σ²⁸-dependent transcription. (A) Efficient stimulation of transcription from the hctB promoter requires the N-terminal residues 1 to 64 and the middle region (residues 114 to 165). One micromolar full-length GrgA (FL) or the indicated deletion mutant was used in each assay. p1 is the P value between basal transcription activity (without GrgA) and activity with GrgA (FL or mutant); p2 is the P value between FL and the deletion mutant (paired t tests of three independent experiments); na, not applicable. (B) Pulldown of NH-GrgA full length (FL) and Δ1-64 but not Δ114-165 by Strep-Tactin bead-immobilized NS-σ²⁸. GrgA and deletion mutants were detected by Western blotting using a mouse anti-GrgA antibody, because the anti-His used in panels A and B does not recognize Δ1-64 (18). (C) Δ138-165 but not Δ114-137 is defective in DNA binding, like Δ114-165. Electrophoresis mobility shift assays (EMSA) were performed using a radiolabeled DNA fragment carrying sequences extending from position −144 to +52 of the defA gene (18) in the presence of the indicated concentrations of wild-type GrgA or the indicated GrgA mutant. (D) Δ138-165 but not Δ114-137 is defective in σ²⁸ binding. Protein pulldown and detection were performed as described for panel B, with the exception of detection via anti-His. (E) Δ138-165 but not Δ114-137 is fully defective in activating transcription from the hctB promoter. See the legend to Fig. 2D for experimental and statistical information.

Our previous studies have shown that deletion of residues 114 to 165 disables GrgA's DNA binding, leading to loss of stimulation of transcription from σ⁶⁶-dependent promoters, whereas removal of residues 1 to 64 disables σ⁶⁶ binding, also causing a defect in activating σ⁶⁶-dependent transcription (18). Therefore, the results shown in Fig. 3A suggest that (i) DNA binding is also required for σ²⁸-dependent transcription and (ii) the N-terminal σ⁶⁶-interacting region interacts with σ²⁸ as well. Surprisingly, pulldown assays demonstrated that GrgAΔ114-165 is completely defective in σ²⁸ binding, whereas GrgAΔ1-64 appeared to have only a slightly decreased σ²⁸-binding activity (Fig. 3B).

We performed a series of deletions within the 114-165 region to define the elements required for interacting with either DNA or σ²⁸. Since residues 114 to 138 are predicted to have coiled and stranded structures, whereas residues 139 to 158 are rich in positively charged lysine and aspartate and are predicted to form a helix (Fig. S4), we expected GrgAΔ114-137 but not GrgAΔ138-165 to retain DNA-binding activity. Electrophoretic mobility shift assay (EMSA) confirmed this prediction (Fig. 3C). Interestingly, GrgAΔ114-137 but not GrgAΔ138-165 also retained σ²⁸-binding activity (Fig. 3D). Not surprisingly, GrgAΔ114-137 but not GrgAΔ138-165 retained the capacity to activate σ²⁸-dependent transcription (Fig. 3E). Additional and extensive deletion mutagenesis and functional analyses for the region of residues 138 to 165 failed to (i) separate residues required for σ²⁸ binding from residues required for DNA binding (Fig. S5 and S6), and (ii) define a smaller region fully required for binding either σ²⁸ or DNA (Fig. S5 and S6). These studies suggest that σ²⁸ and DNA bind to the same region in GrgA and further confirm that σ²⁸ and DNA binding are required for activation of σ²⁸-dependent transcription (Fig. S7).

Residues 1 to 64 in GrgA contribute to σ²⁸ binding.

Transcription assays showed a 50% loss of activity in activating σ²⁸-dependent transcription in the GrgAΔ1-64 mutant (Fig. 3A). We used the BLItz system (20) to confirm decreased σ²⁸-binding activity in Δ1-64. Representative BLItz recordings of binding experiments using full-length NH-GrgA or deletion mutants as ligands and NS-σ²⁸ as analyte are shown in Fig. S8A to D, and kinetic parameters are provided in Table 2. Compared to the full-length NH-GrgA, NS-σ²⁸ revealed a moderately slowed association with and a moderately accelerated dissociation from NH-GrgAΔ1-64, as indicated by a nearly 3-fold decrease in the k_a and a 2-fold increase in the k_d (Table 2). These changes resulted in a highly significant 4.8-fold increase in the K_D value. On the other hand, NH-GrgAΔ114-137, which retains the activity to activate σ²⁸-dependent transcription (Fig. 3E), demonstrated no changes in kinetic parameters for interaction with NS-σ²⁸ (Table 2). In contrast to NH-GrgAΔ114-137, NH-GrgAΔ138-165 immediately and completely dissociated from NS-σ²⁸ upon washing (Fig. S8D), leading to a 1.5 × 10⁶ times higher k_d and a 3.1 × 10⁶ times higher K_D (Table 2), which are fully consistent with pulldown data (Fig. 3D). Furthermore, unlike NS-σ²⁸, which retained a low affinity for NH-GrgAΔ1-64 (relative to full-length NH-GrgA), CS-σ⁶⁶ quickly and completely dissociated from NH-GrgAΔ1-64 upon washing (Fig. S8E), which is consistent with pulldown data previously reported (18). Taken together, the BLItz data shown in Fig. S8 and Table 2 indicate that the decreased affinity with NS-σ²⁸ in NH-GrgAΔ1-64 is responsible for the partial loss of activity in activating σ²⁸-dependent transcription (Fig. 3A).

TABLE 2.

Deletion of amino acids 1 to 64 from GrgA negatively affects σ²⁸ binding^a

Ligand	n	ka		kd		KD
		1/Ms	% control	1/s	% control	M	% control
NH-GrgA	8	(1.5 ± 1.7) × 104	100	(2.8 ± 0.8) × 10−3	100	(2.2 ± 1.1) × 10−7	100
NH-GrgAΔ1-64	4	(5.6 ± 2.8) × 103; P = 0.037	37	(5.8 ± 2.3) × 10−3; P = 0.009	200	(1.1 ± 0.2) × 10−6; P < 0.001	479
NH-GrgAΔ114-137	4	(1.5 ± 0.4) × 104; P = 0.947	98	(3.5 ± 0.5) × 10−3; P = 0.128	124	(2.5 ± 0.3) × 10−7; P = 0.658	112
NH-GrgAΔ138-165	2	(5.6 ± 0.1) × 103; P = 0.125	37	(4.1 ± 0.3) × 102; P = 0.002	1.5 × 108	(6.9 ± 4.5) × 10−2; P < 0.001	3.1 × 108

^aIndicated His-tagged proteins were immobilized on His biosensors as ligands for the NS-σ²⁸ analyte in BLItz assays. Representative graphs of BLItz recordings are shown in Fig. S8. See Table 1 for information regarding kinetic parameters and statistics.

The N terminus of σ²⁸ is most critical for binding GrgA.

We constructed NH-σ²⁸ variants with deletion of the N-terminal leader sequence, σ factor region 2, 3, or 4 (unlike the housekeeping σ factor, σ²⁸, does not contain region 1) (Fig. 4A). All deletion mutants (i.e., NH-σ²⁸ΔNL, NH-σ²⁸ΔR2, NH-σ²⁸ΔR3, and NH-σ²⁸ΔR4) were expressed in Escherichia coli (Fig. 4B). Noticeably, in pulldown assays, NS-GrgA completely failed to pull down the NH-σ²⁸ΔNL and NH-σ²⁸ΔR2 mutants and pulled down only small amounts of NH-σ²⁸ΔR3 and NH-σ²⁸ΔR4 compared to full-length NH-σ²⁸ (Fig. 4C).

FIG 4.

N-terminal leader sequence and region 2 of σ²⁸ are required for interaction with GrgA. (A) Schematic of σ²⁸ and mutants lacking the indicated regions. (B) Western blot showing expression of purified NH-σ²⁸ and mutants. (C) Precipitation of NH-σ²⁸ and mutants by Strep-Tactin-immobilized NS-GrgA. Shown is a Western blot detecting NH-σ²⁸ or mutant. All proteins were resolved via SDS-PAGE and detected using an anti-His antibody.

In BLItz assays, the rate of association with NH-GrgA varied greatly among the σ²⁸ deletion mutants. Whereas NH-σ²⁸ΔNL had essentially the same k_a as full-length NH-σ²⁸, NH-σ²⁸ΔR2 displayed a significant 2-fold reduction in k_a. In contrast, NH-σ²⁸ΔR3 and NH-σ²⁸ΔR4 showed a 3.5-fold increase and a 50% increase in k_a, respectively (Table 3). All mutants demonstrated dramatic increases (10- to 383-fold) in the k_d (Table 3). Consequently, NH-σ²⁸ΔNL and NH-σ²⁸ΔR2 had 345- and 177-fold higher K_D values, respectively, whereas NH-σ²⁸ΔR3 and NH-σ²⁸ΔR4 both demonstrated 7-fold higher K_D values (Table 3). Representative graphs of BLItz recordings are shown in Fig. S9. Taken together, both the pulldown (Fig. 4) and BLI data (Table 3) indicate that the N terminus of σ²⁸ (i.e., NL and R2) interacts with GrgA while R3 and R4 stabilize the GrgA-σ²⁸ binding. Figure 5 summarizes domain structures of GrgA, σ⁶⁶ and σ²⁸, and modes of interaction between GrgA and the σ factors.

TABLE 3.

The N-terminal leader sequence and region 2 of σ²⁸ are required for GrgA binding^a

Analyte	n	ka		kd		KD
		1/Ms	% control	1/s	% control	M	% control
NH-σ28	3	(2.2 ± 0.4) × 104	100	(4.7 ± 0.1) × 10−3	100	(2.2 ± 0.3) × 10−7	100
NH-σ28Δ1-13	3	(2.4 ± 0.4) × 104; P = 0.649	109	(1.8 ± 0.2) × 100; P < 0.001	38,297	(7.6 ± 0.5) × 10−5; P < 0.001	34,545
NH-σ28ΔR2	3	(1.3 ± 0.4) × 104; P = 0.044	59	(5.3 ± 0.3) × 10−1; P = 0.036	11,276	(3.9 ± 1.1) × 10−5; P = 0.003	17,727
NH-σ28ΔR3	3	(8.0 ± 1.6) × 104; P = 0.004	363	(1.2 ± 0.2) × 10−1; P < 0.001	2,553	(1.5 ± 0.2) × 10−6; P < 0.001	714
NH-σ28ΔR4	3	(3.3 ± 0.5) × 104; P = 0.038	150	(4.9 ± 0.3) × 10−2; P = 0.004	1,043	(1.5 ± 0.6) × 10−6; P = 0.018	714

^aBiotinylated NH-GrgA was immobilized on streptavidin biosensors as the ligand for the indicated analytes in BLItz assays. Representative graphs of recordings are shown in Fig. S9. See Table 1 for information regarding kinetic parameters and statistics.

FIG 5.

Domain structures of GrgA, σ⁶⁶ and σ²⁸, and modes of interaction between GrgA and the σ factors. R₁, R₂, R₃, and R₄ are regions conserved across bacterial species. In addition to the nonconserved region (NCR) between R₁ and R₂ in σ⁶⁶, the N-terminal leader sequences, shown in light color, of both σ⁶⁶ and σ²⁸ are also nonconserved. Numbers on GrgA indicate positions of amino acid residues.

DISCUSSION

As an obligate intracellular bacterium, C. trachomatis faces unique challenges at every stage in the developmental cycle (22). Proper expression of certain genes is required for meeting those challenges (11, 12, 23). Surprisingly, only two transcription activators have been identified in C. trachomatis (24). GrgA was first identified as a transcription activator for σ⁶⁶-dependent genes (18). However, the present study has demonstrated that GrgA potentially stimulates expression of σ²⁸-dependent genes as well. Transcription of chlamydial genes is temporally controlled during the developmental cycle (11, 12, 17, 25, 26). Whereas σ⁶⁶ is involved in transcription of most C. trachomatis genes, some late promoters are recognized by σ²⁸ (17). Microarray studies have shown that synthesis of the σ²⁸ mRNA temporally falls behind that of the σ⁶⁶ mRNA (11, 12). Thus, it would be safe to assume that GrgA activates σ⁶⁶-dependent genes in earlier developmental stages. The high concentration of GrgA in the EB (18, 27, 28) also supports the notion that GrgA plays a critical role in activating transcription of immediate-early and early genes, which are dependent on σ⁶⁶ but not σ²⁸. Protein products of these genes, including (but not limited to) TARP (translocated actin recruiting phosphoprotein), Npt1 ATP/ADP translocase, and Inc proteins (inclusion membrane proteins) are critical for cell entry, acquiring energy from the host cell, and establishing and maintaining a niche for chlamydial growth and development inside the inclusion (11, 29, 30). GrgA may also stimulate expression of genes that are important for EB survival. Although the EB was long regarded as a metabolic inert cellular form, active transcription and translation are readily detectable in isolated EBs (14). The presence of GrgA in RBs suggests that GrgA activates midcycle genes that are required for growth and late genes that are needed for conversion of RBs back to EBs. Significantly, we have shown that GrgA activates transcription from σ⁶⁶-dependent early, mid-stage, and late gene promoters in vitro (18).

Whether or not GrgA also regulates expression of σ²⁸-dependent genes during later developmental stages likely depends on the expression levels of GrgA, σ²⁸, and σ⁶⁶. If GrgA is limited, σ²⁸ would have to be present at significantly higher concentrations than σ⁶⁶ to effectively compete for GrgA. However, in a quantitative whole-proteomic mass spectrometry study by Skipp et al., higher levels of GrgA relative to σ⁶⁶ were detected in both EBs and RBs purified from the midcycle, whereas σ²⁸ was undetected in either cellular form (28). Thus, GrgA could stimulate transcription from σ²⁸-dependent promoters in addition to σ⁶⁶-dependent promoters regardless of the molar ratio of the two σ factors. It should be pointed out that another quantitative proteomic study, carried out by Saka at al., failed to detect GrgA in the RBs (27). They also detected a lower concentration of GrgA than σ⁶⁶ (27). Whereas reasons for the discrepancy between the two proteomic studies are unclear, we found that GrgA is readily detectable in RBs (18). Accurate quantification of GrgA and σ factors in different stages of the developmental cycle is needed to elucidate roles of GrgA in the expression of σ⁶⁶- and/or σ²⁸-dependent genes throughout the developmental cycle.

We used both pulldown assays and BLI to analyze the interaction of GrgA with σ²⁸ and σ⁶⁶. Clearly, owing to its quantitative nature, BLI offers higher sensitivities than protein pulldown assays in studying protein-protein interaction. This led to the conclusion that decreased affinity for σ²⁸ in GrgAΔ1-64, which was ambiguous in pulldown assays, is the most probable cause for a 50% loss of activity in activating σ²⁸-dependent transcription.

We have defined a middle region in GrgA (residues 138 to 165) as a σ²⁸- and DNA-binding domain (Fig. 3 and 5). Extensive deletion mutagenesis in this region failed to divide it into subdomains that bind either σ²⁸ or DNA but not both (Fig. S5 and S6). We speculate that multiple positively charged residues (K138, K139, R142, R143, K144, K147, K150, K152, K154-156, R159-161, and/or K164) interact with negatively charged DNA, whereas negatively charged residues (E141, E145, E149, D153, and/or E165) interact with σ²⁸.

GrgA has demonstrated similar but nonidentical properties in activating σ⁶⁶- and σ²⁸-dependent transcription (Fig. 5). Apparently, sequence-nonspecific DNA binding is required for activating both σ⁶⁶-dependent transcription (18) and σ²⁸-dependent transcription (Fig. 3A and E and 5 and Fig. S7). However, the N-terminal region (residues 1 to 64) of GrgA has a stronger role in σ⁶⁶-dependent transcription (18) than in σ²⁸-dependent transcription (Fig. 3A), because this region is absolutely required for GrgA to interact with σ⁶⁶ (18) but plays only a supportive role in binding σ²⁸, which was clearly evident only with BLI (Table 2) but appeared uncertain with pulldown assays (Fig. 3B).

Whereas the major GrgA structural determinants for binding σ²⁸ and σ⁶⁶ differ, there is similarity between the GrgA-binding regions in the two σ factors (Fig. 5). The GrgA-binding sequence in σ⁶⁶ is the last portion of the nonconserved region immediately upstream of conserved region 2, whereas the GrgA-binding sequence in σ²⁸ also involves the N-terminal nonconserved leader sequence (and the immediately downstream region 2). To the best of our knowledge, GrgA is the only transcription factor that targets nonconserved regions of σ factors (16).

In summary, we have demonstrated that the Chlamydia-specific protein GrgA can activate both σ⁶⁶-dependent transcription and σ²⁸-dependent transcription in vitro. Current knowledge suggests that GrgA activates σ⁶⁶-dependent genes throughout the developmental cycle. Whether or not GrgA also regulates expression of σ²⁸-dependent late genes likely depends on the expression levels of GrgA, σ²⁸, and σ⁶⁶, because GrgA has a lower affinity for σ²⁸ than σ⁶⁶. To date, GrgA remains the only transcription factor that targets nonconserved regions of σ factors (16).

MATERIALS AND METHODS

Reagents.

All DNA primers were custom synthesized at Sigma-Aldrich. The QuikChange site-directed mutagenesis kit and BL21(DE3) ArcticExpress E. coli competent cells were purchased from Agilent Technologies. A Q5 site-directed mutagenesis kit and deoxynucleotides were purchased from New England BioLabs. Isopropyl-β-d-thiogalactopyranoside (IPTG) was purchased from Gold Biotechnologies. Talon metal affinity resin was purchased from TaKaRa. The Strep-Tactin Superflow high-capacity resin and d-desthiobiotin were purchased from IBA Life Sciences. Coomassie brilliant blue G-250 dye, mouse monoclonal antihistidine antibody (H1029), goat anti-mouse horseradish peroxidase-conjugated antibody (A4416), and EZ-Link sulfo-NHS-LC-biotin were purchased from Sigma-Aldrich. SuperSignal West Pico Plus chemiluminescent substrate was purchased from ThermoFisher Scientific. Dip and Read nickel-nitrilotriacetic acid (NTA) biosensors were purchased from Pall ForteBio. The HNE buffer contained 50 mM HEPES (pH 7.4), 300 mM NaCl, and 1 mM EDTA. The HNEG buffer contained 50 mM HEPES (pH 7.4), 300 mM NaCl, 1 mM EDTA, and 6 M guanidine HCl. The TNE buffer contained 25 mM Tris (pH 8.0), 150 mM NaCl, and 1 mM EDTA. The protein storage (PS) buffer contained 25 mM Tris-HCl (pH 8.0), 150 mM NaCl, 0.1 mM EDTA, 10 mM MgCl₂, 0.1 mM dithiothreitol (DTT), and 30% (wt/vol) glycerol. The BLItz buffer contained 25 mM Tris-HCl (pH 8.0), 150 mM NaCl, 0.1 mM EDTA, 10 mM MgCl₂, and 0.1 mM DTT.

Vectors.

Plasmids for expressing His- or Strep-tagged GrgA, σ⁶⁶, σ²⁸, and their mutants are listed in the Table S1 in the supplemental material. Sequences of primers used for constructing expression plasmids (GrgA deletion mutants, NS-σ²⁸) and a DNA fragment for EMSAs are provided in Table S2. Sequence authenticities of cloned genes and epitope tags in the final vectors were confirmed using Sanger's DNA sequencing service, provided by GenScript Biotech Corporation.

Expression of recombinant proteins and preparation of cell extract for purification.

BL21(DE3) ArcticExpress E. coli cells transformed with a plasmid for expressing an epitope-tagged chlamydial protein (GrgA, σ²⁸, σ⁶⁶, or their mutant) (Table S1) were cultured in the presence of 1 mM IPTG overnight at 15°C in a shaker. Cells were collected by centrifugation and resuspended in one of the following buffers: HNE buffer (for purification of native His-tagged proteins), HNEG buffer (for purification of denatured His-tagged proteins), or TNE buffer (for purification of native Strep-tagged proteins). The cells were disrupted using a French press. The cell extract was subjected to high-speed (20,000 × g) centrifugation at 4°C for 30 min. Supernatant was collected and used for protein purification.

Purification of Strep-tagged proteins.

Strep-tagged GrgA and σ factors were purified as previously described (19). The supernatant of centrifuged cell lysate was incubated with the Strep-Tactin Superflow high-capacity resin on a Nutator for 1 h at 4°C. The resin was packed onto a column, washed with 30 column volumes of the TNE buffer, and then eluted with the TNE buffer containing 2.5 mM d-desthiobiotin. The elution was collected in 10 fractions. Protein in the fractions was examined following SDS-PAGE and Coomassie blue staining. Fractions with high purity and concentration were pooled and dialyzed overnight against the PS buffer at 4°C and then stored in aliquots at −80°C.

Purification of His-tagged proteins.

The supernatant of centrifuged cell lysate was incubated with the Talon metal affinity resin on a Nutator for 1 h at 4°C. The resin incubated with nondenatured cell extract was packed onto a column, washed with 30 column volumes of HNE buffer containing 1% NP-40, and eluted with the HNE buffer containing 250 mM imidazole. The resin incubated with denatured cell extract was packed onto a column, washed with 30 column volumes of HNEG buffer, and eluted with HNEG buffer containing 250 mM imidazole. Examination of protein purity, dialysis, and storage was carried out in the same manner as that for purified Strep-tagged proteins (18).

In vitro transcription assay.

In vitro transcription of σ²⁸-dependent promoter was performed as previously described (18). The assay, in a total volume of 30 μl, contained 200 ng supercoiled plasmid DNA, 50 mM potassium acetate, 8.1 mM magnesium acetate, 50 mM Tris acetate (pH 8.0), 27 mM ammonium acetate, 1 mM DTT, 3.5% (wt/vol) polyethylene glycol (average molecular weight, 8,000), 330 μM ATP, 330 μM UTP, 1 μM CTP, 0.2 μM [α-³²P]CTP (3,000 Ci/mmol), 100 μM 3′-O-methyl-GTP, 20 U of RNasin, RNAP, and the indicated amount of GrgA or GrgA mutant. The reactions using cRNAP and σ²⁸ contained 1.0 μl purified cRNAP and 30 nM His-tagged σ²⁸, purified by procedures involving denaturing and refolding as described above. For reactions using eCore and σ²⁸, their concentrations were 5 nM and 30 nM, respectively. The reaction was allowed to continue at 37°C for 40 min and terminated by the addition of 70 μl of 2.86 M ammonium acetate containing 4 mg of glycogen. After ethanol precipitation, ³²P-labeled RNA was resolved by 8 M urea–6% polyacrylamide gel electrophoresis and quantified with a Storm PhosphorImager and ImageQuant software. Relative amounts of transcripts were presented, with that of the control reaction set as 1 U. Data shown in bar graphs represent averages ± standard deviations (SDs) from three or more independent experiments. Pairwise, two-tailed Student t tests were used to compare data.

EMSA.

GrgA-DNA interaction was determined by EMSA as described previously (18). ³²P-labeled DNA fragment containing the C. trachomatis defA promoter (31) was amplified using a ³²P-labeled 5′ primer and an unlabeled 3′ primer (Table S2) and purified with a Qiagen column. The GrgA-DNA binding reaction was performed in a total volume of 10 μl, containing 10 nM promoter fragment, the indicated amount of NH-GrgA, 1 mM potassium acetate, 8.1 mM magnesium acetate, 50 mM Tris acetate (pH 8.0), 27 mM ammonium acetate, 1 mM DTT, and 3.5% (wt/vol) polyethylene glycol (average molecular weight, 8,000). After mixing for 1 h at 4°C, the binding mixture was resolved by 6% nondenaturing polyacrylamide gel. Free and GrgA-bound DNA fragments were visualized on a Storm PhosphorImager (Molecular Dynamics).

Pulldown assays.

Twenty microliters of Strep-Tactin Superflow high-capacity resin was washed twice with the HNE buffer and incubated with 50 μl of Strep-tagged cell extract or purified protein on a Nutator at 4°C for 1 h. The resin was washed three times with HNE buffer containing 1% NP-40 and then incubated with 5 μg of a purified His-tagged protein (or mutant) on a Nutator at 4°C for 1 h. After 3 washes with the HNE buffer containing 1% NP-40 and a final wash with phosphate-buffered saline (PBS), the resin was eluted using SDS-PAGE sample buffer. All protein was resolved via SDS-PAGE and detected by either Coomassie blue staining or Western blotting using a monoclonal mouse anti-His or a polyclonal mouse anti-GrgA primary antibody and horseradish peroxidase-conjugated goat anti-mouse secondary antibody.

Preparation of biotinylated protein.

Purified NH-GrgA was dialyzed against PBS to remove Tris and then incubated with 10 mM EZ-Link sulfo-NHS-LC-biotin for 2 h at 4°C. Excess biotin was removed via two-step dialysis, initially against PBS and subsequently against the PS buffer.

BLI assay.

An NTA His or streptavidin biosensor was subjected to initial hydration in BLItz buffer for 10 min before being loaded onto the ForteBio BLItz machine and washed with BLItz buffer for 30 s to obtain a baseline reading. The His biosensor was then incubated with 4 μl of a His-tagged ligand for 240 s. The concentrations of ligand ranged from 1 to 20 μM, all of which saturated the His-binding sites on the biosensor. Alternatively, the streptavidin biosensor was incubated with 4 μl of a biotinylated ligand (10 μM NH-GrgA, which was sufficient to saturate the binding sites on the biosensor) for 240 s. After a brief wash with BLItz buffer for 30 s to remove excess protein, the biosensor was incubated with 4 μl of an analyte (purified Strep-tagged protein for the His biosensor or NH-σ²⁸ for the streptavidin biosensor) for 120 s to measure association of the ligand-analyte complex. Subsequently, the biosensor was washed with BLItz buffer for 120 s to measure disassociation of the ligand-analyte complex. All BLItz recordings were subsequently fit to a 1:1 binding model using BLItz Pro software (version 1.1.0.31), which generated the association rate constant (k_a), disassociation rate constant (k_d), and disassociation equilibrium constant (K_D) for each interaction.