ABSTRACT
The sexually transmitted 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 non-conserved 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 σ66and 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 a NCR located upstream of conserved region 2 of σ28. Our findings suggest 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 under-developed countries. The pathogen has a unique developmental cycle, which is transcriptionally regulated. The discovery of an expanded role for the Chlamydia-specific transcription factor GrgA helps understand progression of the chlamydial developmental cycle.
INTRODUCTION
Each year, about 2.2 million cases of notifiable infections are reported to the Centers for Disease Control and Prevention (CDC). These infections are caused by nearly 100 different pathogens, but the majority (about 1.6 million, i.e., 60%) is due to the sexually transmitted pathogen Chlamydia trachomatis (1, 2). Still, CDC estimates that only 1 tenth of C. trachomatis-infected cases are reported because the infection is mostly asymptomatic (3). Nonetheless, without proper antibiotic treatment, the infection often leads to serious complications. In fact, C. trachomatis is the most common infectious cause of infertility and pelvic inflammatory syndrome in women. Infection in pregnant women may result in abortion or premature birth. Pathological changes in the fallopian tubes caused by C. trachomatis infection may lead to ectopic pregnancy, which causes severe bleeding and likely death if the ectopically embedded embryo is not detected and terminated early enough. Infants may develop C. trachomatis pneumonia following acquisition of the pathogen while passing the birth canal of an infected mother. 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 intracellularGram-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 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 h. No longer infectious, the RB divides exponentially by binary fission until around 20 h when a significant portion of RBs re-differentiate back into EBs while some RBs continue proliferation (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 encodes fewer than 1000 genes (10). Microarray analyses demonstrated that the majority of these genes are transcribed starting a few hours post-inoculation 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). 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 α 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 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 the gene codes CT_504 and CTL0766 for C. trachomatis serovar 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 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 pull-down assays using differential epitope-tagging. The StrepTactin 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.
GrgA has a lower affinity for σ28 than for σ66
Next, we 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 pull-down assays. As expected, NS-GrgA efficiently pulled down NH-σ28 and CH-σ66 in separate reactions (Fig. S1). 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 next quantitatively characterized GrgA-binding by σ28 and σ66 with biolayer interferometry 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 CS-σ66 and NS-σ28 an analyte are shown in Fig. S2A, B), values of kinetic parameters are provided in Table 1. The CS-σ66 analyte yielded a statistically highly significant 25-fold higher ka than 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 kd, 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 KD, indicating that GrgA has a lower overall affinity for σ28 than σ66.
Reciprocal BLI using CH-σ66 and NH-σ28 as ligands and NS-GrgA as the analyte were performed to validate the difference in GrgA binding by the σ factors presented above (Fig. S2C, D and Table 1). Consistent with the trend in ka value changes presented above, the NS-GrgA analyte also demonstrated a statistically significant higher ka for CH-σ66 than for NH-σ28 although the difference is smaller (25-fold vs 3.7 fold). Interestingly, the kd values reveal that NS-GrgA also dissociates from CH-σ66 6-fold slower than from NH-σ28. Compared to the CH-σ66-NS-GrgA interaction, the NH-σ28-NS-GrgA interaction had a 28fold higher KD, which is nearly identical to the 32-fold higher KD detected for the NH-GrgA-NS-σ28 interaction vs the NH-GrgA-CS-σ66 interaction. Thus, competitive pull-down 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 (Fig. 2A). Interestingly, GrgA demonstrated a dose-dependent stimulatory effect on the transcription from the promoter (Fig. 2B, 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.
Residues 138-165 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, B) were tested for the effects on transcription from the hctB promoter (Fig. 3A). Noticeably, GrgAΔ 114-165 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 (65-113, 166266 and 207-288) from GrgA had either no or minimal effects on the transcription activation (Fig. 3A).
Our previous studies have shown that deletion of residues 114-165 disables GrgA’s DNA-binding, leading to loss of stimulation of transcription from σ66-dependent promoters, whereas removal of residues 1-64 disables σ66-binding, also causing defect in activating σ66-dependent transcription (18). Therefore, the results in Fig. 3A suggests that 1) DNA-binding is also required for σ28-dependent transcription, and 2) the N-terminal σ66-interacting region may interact with σ28 as well. Surprisingly, pull-down 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 elementsrequired for interacting with either DNA or σ28. Since residues 114-138 are predicted to have coiled and stranded structures, whereas residues 139-158 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. EMSA confirmed this prediction (Fig. 3C). Interestingly, GrgAΔ114-137 but not GrgAΔ138-165 also retained σ28-binding activity as well (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 138-165 failed to 1) separate residues required for σ28-binding from residues required for DNA-binding (Fig. S5 & S6), and 2) define a smaller region fully required for binding either σ28 or DNA (Fig. S5 & 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 1-64 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-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 the NH-GrgAΔ1-64, as indicated by a nearly 3-fold decrease in the ka and a 2-fold increase in the kd (Table 2). These changes resulted in a highly significant 4.8-fold increase in the KD 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 wash (Fig. S8D), leading to a 1.5 X 106 times higher kd and a 3.1 X 106 times higher KD (Table 2), which are fully consistent with pull-down 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 wash (Fig. S8E), which is consistent with pull-down data previously reported (18). Taken together, the BLItz data 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).
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 E. coli (Fig. 4B). Noticeably, in pull-down 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).
In BLItz assays, the rate of association with NH-GrgA varied greatly among the σ28 deletion mutants. Whereas NH-σ28ΔNL had essentially the same ka as full length NH-σ28, NH-σ28ΔR2 displayed a significant 2-fold reduction in ka. In contrast, NH-σ28ΔR3 and NH-σ28ΔR4 showed a 3.5-fold increase and a 50% increase in ka, respectively (Table 3). All mutants demonstrated dramatic increases (10-383 fold) in the kd (Table 3). Consequently, NH-σ28ΔNL and NH-σ28ΔR2 had 345- and 177-fold higher kD value, respectively, whereas NH-σ28ΔR3 and NH-σ28ΔR4 both demonstrated 7 fold higher KD values (Table 3). Representative graphs of BLItz recordings are shown in Fig. S9. Taken together, both the pull-down (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.
DISCUSSION
Although GrgA was first identified as a transcription activator for σ66-dependent genes (18), the present study has demonstrated that GrgA potentially stimulates expression of σ28-dependent genes. Transcription of chlamydial genes is temporally controlled during the developmental cycle (11, 12, 17, 22, 23). 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 the σ66 mRNA (11, 12). Thus, it would be safe to assume that GrgA primarily activates σ66-dependent genes in earlier developmental stages.
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, quantitative whole proteomic mass spectrometry analyses detected higher levels of GrgA relative to σ66 in both EBs and RBs purified from the midcycle (24) whereas σ28 was undetected in either cellular form (24, 25). Thus, GrgA could potentially stimulate transcription from σ28-dependent promoters in addition to σ66- dependent promoters regardless the molar ratio of the two σ factors. Accurate quantification of GrgA and a factors in different stages of the developmental cycle will help elucidate the role of GrgA in the expression of σ66- and/or σ28-dependent genes in different developmental stages.
We used both pull-down 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 pull-down assays in studying protein-protein interaction. This led to the confirmation that decreased affinity for σ28 in GrgAΔ1-64, which was ambiguous in pull-down 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 138-165) as a σ28- and DNA-binding domain (Fig. 3). Extensive deletion mutagenesis in this region failed to divide it into subdomains that bind either σ28 or DNA but not both (Figs. S5 & 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 multiple negatively-charged residues E141, E145, E149, D153 and/or E165) interact with σ28.
GrgA has demonstrated similar but not identical properties in activating σ66- and σ28-dependent transcription. Apparently, sequence-nonspecific DNA-binding is required for activating both σ66- dependent transcription (18) and σ28-dependent transcription (Fig. 3A, E & Fig. S7). However, the N-terminal region (residues 1-64) 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 pull-down 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 a factors. The GrgA-binding sequence in σ66 is the last portion of the non-conserved region immediately upstream of the conserved region 2, whereas the GrgA-binding sequence in σ28 also involves the N-terminal non-conserved leader sequence (and the immediately downstream region 2). To the best of our knowledge, GrgA is the only transcription factor that targets non-conserved regions of σ factors (16).
In summary, we have demonstrated that the Chlamydia-specific GrgA can activate both σ66-dependent transcription and σ28-dependent transcription in vitro. Current knowledge suggests that GrgA primarily activates σ66-dependent genes during earlier developmental stages. However, 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 because GrgA has a lower affinity for σ28 than σ66. To date, GrgA remains the only transcription factor that targets non-conserved regions of σ factors (16).
MATERIALS AND METHODS
Reagents
All DNA primers were custom-synthesized at Sigma Aldrich. The QuikChange Site-Directed Mutagenesis Kit, BL21(DE3) ArcticExpress E. coli competent cells were purchased from Agilent Technologies. Q5 Site-Directed Mutagenesis Kit, and deoxynucleotides were purchased from New England BioLabs. Isopropyl β-D-1-thiogalactopyranoside (IPTG) was purchased from Gold Biotechnologies. TALON Metal Affinity Resin was purchased from Takara. The StrepTactin superflow high capacity resin and D-desthiobiotin were purchased from IBA Life Sciences. Coomassie Brilliant Blue G-250 Dye, mouse monoclonal anti-Histidine antibody (H1029), goat anti-mouse horseradish-perodixase-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 Ni-NTA (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 6M 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 DTT, and 30% glycerol (w/v). The BLItz Buffer contained 25 mMTris-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. Sequences of primers used for constructing expression plasmids (GrgA deletion mutants, NS-σ28)and a DNA fragment for EMSA assays are available upon request. 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 (20,000g) centrifugation at 4 °C for 30 minutes. 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 StrepTactin superflow high capacity resin on a Nutator for 1 h at 4 °C. The resin was packed onto a column and 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 hour at 4 °C. The resin incubated with non-denatured 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 were carried out in the same manner as for purified Strep-tagged proteins (18).
In vitro transcription assay
In vitro transcription of σ28-dependent promoter was performed as previously described (18). Theassay in a total volume of 30 μl contained 200 ng supercoiled plasmid DNA, 50 mM potassium acetate, 8.1mM magnesium acetate, 50 mM Tris acetate (pH8.0), 27 mM ammonium acetate, 1 mM DTT, 3.5% (wt/vol) poly-ethylene 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 units of RNasin, RNAP, and indicated amount of GrgA or GrgA mutant. The reactions using cRNAP and σ28 contained 1.0 μL purified cRNAP and 30nM 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 pursue at 37 °C for 40 min and terminated by the addition of 70 μL of 2.86 M ammonium acetate containing 4mg of glycogen. After ethanol precipitation, 32P-labeled RNA was resolved by 8M urea-6% polyacrylamide gel electrophoresis, and quantified with a Storm Phosphorimager and the ImageQuant software. Relative amounts of transcripts were presented with that of the control reaction set as 1 unit. Data shown in bar graphs represent averages ± SDs from three or more independent experiments. Pairwise, two-tailed Student t tests were used to compare data.
Electrophoresis mobility shift assay (EMSA)
GrgA-DNA interaction was determined by EMSA as described previously (18). 32P-labeled DNA fragment containing the C. trachomatis defA promoter (26) 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, an 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% non-denaturing polyacrylamide gel. Free and GrgA-bound DNA fragments were visualized on a Storm Phosphorimager (Molecular Dynamics).
Pull-down assays
20 μL of StrepTactin 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 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 HRP-conjugated goat anti-mouse secondary antibody.
Preparation of biotinylatedprotein
Purified NH-GrgA was dialyzed against PBS to remove Tris and then incubated with 10 mM EZ-Link Sulfo-NHS-LC-Biotin for 2 hours at 4 °C. Excess biotin was removed via two-step dialysis, initially against PBS and subsequently against the PS buffer.
Bio-layer interferometry assay
An NTA His or streptavidin biosensor was subjected to initial hydration in BLItz Buffer for 10 minutes before being loaded onto the ForteBio BLItz machine and washed with BLItz Buffer for 30 seconds to obtain a baseline reading. The His biosensor was then incubated with 4 μL of a His-tagged ligand for 240 seconds. The concentration of ligand ranged from 1-20 μM, which all saturated the His-binding sites on the biosensor. Alternatively, the streptavidin biosensor was incubated with 4 μL of a biotinylated ligand (NH-GrgA, 10 μM, which was sufficient to saturate the binding sites on the biosensor) for 240 seconds. After a brief wash with BLItz Buffer for 30 seconds 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 seconds to measure association of the ligand-analyte complex. Subsequently, the biosensor was washed with BLItz Buffer for 120 seconds to measure disassociation of the ligand-analyte complex. All BLItz recordings were subsequently fit to a 1:1 binding model using the BLItz Pro software (version 1.1.0.31), which generated the association rate constant (ka), disassociation rate constant (kd), and disassociation equilibrium constant (KD) for each interaction.
ACKNOWLEDGEMENTS
We thank Ming Tan (University of California Irvine) for pMT1212. This research was supported by National Institutes of Health (Grant # AI122034 to HF, GM118059 to BEN), New Jersey Health Foundation (Grant # PC 20-18 to HF) and Natural Sciences Foundation of China (Grant # 31370209 and 31400165 to XB).