Repurposing HDAC inhibitors to enhance ribonuclease 4 and 7 expression and reduce urinary tract infection

Significance

With the emergence of antibiotic-resistant bacteria, new approaches are needed for the treatment of urinary tract infections. Host defense peptides have desirable features as antimicrobials and may provide an alternative to antibiotics. Ribonuclease 4 and ribonuclease 7 are host defense peptides produced by the kidney and bladder and exhibit bactericidal activity against uropathogenic bacteria and antibiotic-resistant bacteria. Identifying ways to boost their expression may represent a unique approach to treat urinary tract infections and conserve antibiotic use.

Abstract

With the emergence of antibiotic-resistant bacteria, innovative approaches are needed for the treatment of urinary tract infections. Boosting antimicrobial peptide expression may provide an alternative to antibiotics. Here, we developed reporter cell lines and performed a high-throughput screen of clinically used drugs to identify compounds that boost ribonuclease 4 and 7 expression (RNase 4 and 7), peptides that have antimicrobial activity against antibiotic-resistant uropathogens. This screen identified histone deacetylase (HDAC) inhibitors as effective RNase 4 and RNase 7 inducers. Validation studies in primary human kidney and bladder cells confirmed pan-HDAC inhibitors as well as the HDAC class I inhibitor, MS-275, induce RNase 4 and RNase 7 to protect human kidney and bladder cells from uropathogenic Escherichia coli. When we administered MS-275 to mice, RNase 4 and 7 expression increased and mice were protected from acute transurethral E. coli challenge. In support of this mechanism, MS-275 treatment increased acetylated histone H3 binding to the RNASE4 and RNASE7 promoters. Overexpression and knockdown of HDAC class I proteins identified HDAC3 as a primary regulator of RNase 4 and 7. These results demonstrate the protective effects of enhancing RNase 4 and RNase 7, opening the door to repurposing medications as antibiotic conserving therapeutics for urinary tract infection.

Urinary tract infection (UTI) is one of the most common bacterial infections, impacting more than 150 million people annually. In the United States, UTI accounts for over 8.6 million annual office visits, 600,000 hospitalizations, and $4 billion in hospital costs (1, 2). UTIs are a significant cause of morbidity in infant boys, older men, and women of all ages. Half of all women experience a UTI by 30 y of age and approximately 30% of women develop recurrent infections. UTI sequelae include recurrent infections, pyelonephritis with kidney scarring, urosepsis, and preterm birth. A range of pathogens have been identified to cause UTI but uropathogenic Escherichia coli (UPEC) are identified as the inciting pathogen in more than 80% of cases (3). Antibiotics are the cornerstone of UTI treatment. However, antibiotic overuse has contributed to the development and worldwide dissemination of antibiotic-resistant uropathogens, reducing our capacity to eliminate this common infection. Concurrently, the pipeline of new antimicrobials in development has not matched the pace of antibiotic resistance in uropathogenic bacteria. As a result, new UTI prevention and treatment strategies are needed that are not reliant on antibiotics (4, 5).

Recent interest in the molecular defenses of the kidney and bladder suggests that antimicrobial peptides (AMPs) have the potential to be developed as UTI therapeutics (6, 7). AMPs are small cationic peptides that are produced and secreted by cells involved in host defense. In the urinary tract, AMPs are produced by the bladder urothelium and the kidney’s tubules. Certain AMPs exhibit a narrow antimicrobial spectrum, while others show a broad-spectrum antimicrobial activity. AMPs that have been shown to prevent UTI include defensins, cathelicidin, lectins, metal binding proteins, and peptides in the ribonuclease (RNase) A superfamily (8–18). Identifying mechanisms that regulate AMP expression may reveal new UTI treatments that expand therapies beyond the scope of antibiotics.

Our research team has demonstrated that RNase A Superfamily members contribute to UTI defense (8–13). Human and mouse RNase 4 as well as human RNase 7 are produced by the bladder urothelium and kidney collecting duct, which contain cells targeted by UPEC. RNase 4 and RNase 7 have antimicrobial activity against UPEC and antibiotic-resistant UPEC (9, 13). Their catalytic activity is not required for their bactericidal activity. Instead, their antibacterial functions are dependent on their binding affinity to bacterial cell wall proteins, including lipopolysaccharides on gram-negative bacteria, as well as their capacity to disrupt and permeate bacterial membranes. These mechanisms of bacterial membrane disruption differ from antibiotics, which kill or prevent bacterial replication by inhibiting cell wall synthesis, DNA replication, RNA transcription, or protein synthesis (13, 19, 20). Gram-negative bacteria employ several mechanisms to resist killing by AMPs. They express efflux pumps, produce capsule polysaccharides that shield their cell surface, covalently modify their lipopolysaccharide to reduce AMP binding, produce proteases to cleave AMPs, or downregulate AMP expression by host cells (21–23). If RNase 4 or RNase 7 expression is downregulated or silenced in vitro, UPEC attachment to and invasion of bladder urothelial cells increases. In contrast, when RNase 4 and RNase 7 are induced or overexpressed, the ability of UPEC to cause invasive infection decreases (9, 12). These findings suggest that identifying ways to enhance RNase 4 and RNase 7 expression may reduce UTI risk.

Here, we perform a high-throughput screen of medications approved by the United States Food and Drug Administration (FDA) to detect inducers of RNase 4 and RNase 7 expression. In doing so, we identify histone deacetylase inhibitors (HDACi) as a class of drugs that enhance RNase 4 and RNase 7 expression—providing insight into new mechanisms that may be developed as antibiotic-conserving UTI therapies.

Results

Development and Validation of RNASE4 and RNASE7 Luciferase Reporter Assays.

To screen candidate compounds that enhance RNASE4 and RNASE7 expression, we generated human embryonic kidney (HEK)-293 cells stably expressing the luc2 synthetic firefly luciferase gene under the control of the human RNASE4 (HEK293-R4-Luc) or RNASE7 (HEK293-R7-Luc) promoters. With this system, cellular firefly luciferase expression increases when the RNASE4 or RNASE7 promoters are activated (SI Appendix, Fig. S1A). To determine whether HEK293-R4-Luc and HEK293-R7-Luc cells are suitable for high-throughput drug screening, we stimulated them with phorbol myristate acetate (PMA; positive control) or vehicle (dimethyl sulfoxide, DMSO; negative control) for 6 h. Following PMA treatment, we calculated their Z’-factor, a dimensionless screening window coefficient used to quantify the suitability of an assay for a full-scale, high-throughput screen. The Z’-factor measures an assay’s dynamic range or separation between positive and negative controls. HEK293-R4-Luc cells achieved a Z’-factor of 0.62, and HEK293-R7-Luc cells achieved a Z’-factor of 0.75 (SI Appendix, Fig. S1 B and C). Z’-factor values between 0.5 and 1 predict good performance in high-throughput screenings, confirming that our cell-based reporter assays are suitable for high-throughput screenings (24).

Screening an FDA-Approved Drug Library Demonstrates Histone Deacetylase Inhibitors Increase RNASE4 and RNASE7 Promoter Activity.

Using these HEK293-R4-Luc and HEK293-R7-Luc reporter cells, we screened a drug library containing 1,280 compounds (Fig. 1A and Dataset S1). High-throughput screening identified 30 drugs that induce RNASE4 promoter activity and 64 drugs that induce RNASE7 promoter activity at least 2 SDs above vehicle. No compounds were found to suppress the RNASE4 promoter activity, while five compounds suppressed RNASE7 promoter activity 2 SDs below vehicle (Fig. 1 B and G).

Fig. 1.

Drug repurposing library screen identifies histone deacetylase inhibitors as inducers of RNASE4 and RNASE7 promoter activity. (A) Schematic of the experimental design using plasmids expressing firefly luciferase under control of the human RNASE4 or RNASE7 promoters (Left). These plasmids were stably transfected into HEK293 cells to establish HEK293-R4-Luc and HEK293-R7-Luc reporter cells (Center). A high-throughput drug screen was performed with measurement of luciferase activity (luminescence) as the final readout (Right). (B and G) Z-score scatter plots of 1,280 tested drugs were calculated as described in the methods for HEK293-R4-Luc (B) and HEK293-R7-Luc (G) reporter cells. Each point denotes the Z-score of a unique compound. Drugs achieving a |Z-score| greater than or equal to two were considered positive or negative hits (above or below the dotted red line). Z-scores for each compound for either Hek293-R4-Luc or Hek293-R7-Luc are listed in Dataset S1 (C and H) Fold change in luciferase activity was calculated for the top 10 inducers of HEK293-R4-Luc (C) or HEK293-R7-Luc (H) promoter activity compared to vehicle. HDAC inhibitors are identified by the gray bars. (D–F and I–K) Luciferase activity was measured in HEK293-R4-Luc (D–F) or HEK293-R7-Luc (I–K) cells treated with increasing concentrations of Vorinostat, Givinostat, or MS-275. Graphs show the mean and SEM fold induction above vehicle-treated cells. Results are from three independent experiments performed in triplicate (n = 3). Asterisks denote significant P values normalized to vehicle as determined by one-way ANOVA. *P < 0.05, **P < 0.01, ***P < 0.001.

Confirmation assays were performed, as defined in the methods, with drugs from the initial screen that were identified as RNASE4 and RNASE7 inducers. When delineating the compounds that induce the greatest fold change in RNASE4 promoter activity, 9 of the top 10 drugs were pan-histone deacetylase inhibitors (HDACi)—drugs that inhibit class I, IIa, IIb, and IV histone deacetylases (HDAC). Similarly, seven of the top 10 candidates enhancing RNASE7 promoter activity were pan-HDACi (Fig. 1 C and H). These high-throughput screening results suggest HDAC inhibition augments RNASE4 and RNASE7 promoter activity.

Pan-HDAC Inhibitors and Class-Selective HDAC Inhibitors Augment RNASE4 and RNASE7 Promoter Activity.

To validate that HDAC inhibition triggers RNASE4 and RNASE7 promoter activity, we treated HEK293-R4-Luc and HEK293-R7-Luc reporter cells with increasing concentrations of different pan-HDACi for six or 24 h—including Vorinostat, AR-42, Belinostat, Dacinostat, and Quisinostat. Twenty-four hours after treatment, these pan-HDACi maximally induced luciferase activity in HEK293-R4-Luc and HEK293-R7-Luc cells (Fig. 1 D and I and SI Appendix, Fig. S2), confirming the high-throughput screening findings that pan-HDACi enhance RNASE4 and RNASE7 promoter activity.

Next, we treated HEK293-R4-Luc and HEK293-R7-Luc reporter cells with a series of HDACi that have increasing selectivity to identify which HDAC class(es) enhance RNASE4 and RNASE7 promoter activity (SI Appendix, Table S1). Givinostat, which inhibits HDAC class I, IIa, and IIb, augmented RNASE4 and RNASE7 promoter activity 6 and 24 h post treatment (Fig. 1 E and J). Similarly, the more selective HDAC class I inhibitor, MS-275 (Entinostat), enhanced RNASE4 and RNASE7 promoter activity in a dose-dependent manner (Fig. 1 F and K). In contrast, the HDAC class III inhibitor, nicotinamide, did not induce RNASE4 or RNASE7 promoter activity (SI Appendix, Fig. S3). These findings suggest that class I, II, and IV HDAC proteins regulate RNASE4 and RNASE7, while class III HDAC proteins may not have as significant a role.

Histone Deacetylase Inhibitors Induce RNase 4 and RNase 7 Expression in Human Bladder and Kidney Cells.

To evaluate the effects of HDACi on endogenous RNase 4 and RNase 7 expression, we treated primary human bladder urothelial cells (HUC) and human kidney epithelial cells (HKC) with Givinostat or the more selective MS-275. During these experiments, Givinostat or MS-275 treatment did not compromise cell integrity as assessed by lactate dehydrogenase release or trypan blue exclusion assays (SI Appendix, Fig. S4). Quantitative real-time PCR (qRT-PCR) shows Givinostat and MS-275 induced RNASE4 and RNASE7 expression (Fig. 2 A and D). These HDACi also augmented the transcription of other AMPs expressed in human bladder and kidney cells—including CAMP, DEFB1, and LCN2 (SI Appendix, Fig. S5).

Fig. 2.

Givinostat and MS-275 induce RNase 4 and RNase 7 expression in human kidney and bladder cells. (A and D) Relative RNASE4 and RNASE7 transcript expression in human kidney epithelial cells (A) or bladder urothelial cells (D) treated with Givinostat or MS-275 for 24 h. RNASE4 and RNASE7 expression were calculated from five or six independent experiments performed in triplicate (n = 5 to 6). Graphs show the mean transcript expression and SEM. (B and E) Representative Western blots probed for RNase 4, RNase 7, acetyl-Histone H3 (lysine 27), and GAPDH from human kidney epithelial cells (B) or bladder urothelial cells (E) following Givinostat or MS-275 treatment for 24 h. RNase 7 was not routinely detected by Western blot in kidney epithelial cells (B). (C and F) ELISA quantified RNase 4 and RNase 7 concentrations in media isolated from vehicle- or HDACi-treated human kidney epithelial cells (C) or bladder urothelial cells (F) following 24 h treatment with Givinostat or MS-275. Graphs show mean concentrations and SEM. Asterisks indicate significant P values compared to vehicle as determined by one-way ANOVA. *P < 0.05, **P < 0.01, ***P < 0.001.

To further characterize how HDAC inhibition affects RNase 4 and RNase 7 expression, we investigated the cellular production and secretion of RNase 4 and RNase 7 peptides. In HKCs, Western blot demonstrates that Givinostat and MS-275 induce RNase 4 expression. RNase 7 was not routinely detected in these cells by Western blot (Fig. 2B). In HUCs, Western blot shows that Givinostat and MS-275 induce RNase 4 and RNase 7 production (Fig. 2E). In HKCs and HUCs, we quantified RNase 4 and RNase 7 secretion into culture media via ELISA. Twenty-four hours after HKCs were treated with HDACi, Givinostat increased RNase 4 and RNase 7 media concentrations 2.3- and 5.3-fold, respectively. Similarly, MS-275 increased RNase 4 and RNase 7 media concentrations 2.6- and 3.9-fold (Fig. 2C). In HUCs, RNase 4 and Rnase 7 media concentrations increased 2.6- and 3.6-fold following Givinostat treatment while RNase 4 and RNase 7 media concentrations increased 3.1- and 3.5-fold following MS-275 treatment (Fig. 2F). Collectively, these results demonstrate that Givinostat and MS-275 stimulate the transcription of AMP genes as well as the cellular production and secretion of RNase 4 and RNase 7.

HDAC Inhibition Increases Epithelial Defense against Uropathogenic E. coli In Vitro.

To test whether HDAC inhibition enhances immune defenses to shield the kidney epithelia from bacterial challenge, we isolated culture media from HKCs treated with vehicle, Givinostat, or MS-275 and inoculated it with UPEC. UPEC survival decreased in media isolated from Givinostat- or MS-275-treated cells (Fig. 3A). To determine whether HDAC inhibition heightens cellular resistance to bacterial infection, we treated confluent HKCs with Givinostat or MS-275 and challenged them with UPEC. Following bacterial challenge, the percentage of UPEC binding to or invading Givinostat- or MS-275-treated cells was significantly reduced compared to that of vehicle-treated cells (Fig. 3 B and C). These findings indicate that HDACi augment the extracellular and cellular antimicrobial capacity of HKCs to reduce their susceptibility to UPEC.

Fig. 3.

Givinostat and MS-275 protect human kidney and bladder cells from uropathogenic E. coli by boosting RNase 4 and RNase 7 expression. (A) Isolated conditioned culture media from vehicle or HDACi-treated human kidney epithelial cells was inoculated with UPEC (strain CFT073). The number of UPEC were enumerated after a 90-min incubation. Results are from eight independent experiments performed in duplicate (n = 8). Data show the mean UPEC colony forming units (CFU)/mL and SEM. (B and C) Human kidney epithelial cells were treated with Givinostat (500 nM) or MS-275 (500 nM) and challenged with UPEC (strain CFT073). Shown are the percentage of bacteria adhering to the cellular surface (Left) or invading the cells (Right). Graphs show the mean and SEM. Results, are from six independent experiments performed in triplicate (n = 6). (D) Primary human bladder urothelial cells were transiently transfected with an RNASE4 RNASE7, or a nontargeting control (NTC) control siRNA pool and treated with MS-275 (500 nM). qRT-PCR confirmed RNASE4 (Left) and RNASE7 knockdown (Right). (E) Extracellular UPEC kill assays were performed in culture media isolated from vehicle- or MS-275-treated human urothelial cells. Graphs show mean UPEC survival and SEM. Results are from five independent experiments performed in triplicate (n = 5). (F) Following RNASE4 and RNASE7 silencing, UPEC attachment and invasion assays were performed on primary human urothelial cells treated with MS-275 (500 nM). Shown are the percentage of UPEC (strain UTI89) adhering to the cellular surface (Left) or invading the cells (Right). Graphs show the mean and SEM. Results are from five or six independent experiments performed in triplicate (n = 5 to 6). Asterisks indicate significant P values for the pairwise comparisons as determined by one-way ANOVA (A and D–F) or Student’s t test (B and C). *P < 0.05, **P < 0.01, ***P < 0.001.

To define the contributions of RNase 4 and RNase 7 to cellular defense against UPEC, HUCs were transiently transfected with an RNASE4, RNASE7, or nontargeting control small interfering RNA (siRNA) pool. Three days after transfection, cells were treated with MS-275. qRT-PCR verified RNASE4 and RNASE7 knockdown (Fig. 3D). Media assays confirmed UPEC survival decreased in conditioned media isolated from MS-275-treated HUCs. This effect was mitigated when RNASE4 or RNASE7 was silenced (Fig. 3E). Additionally, UPEC attachment to the cellular surface and cellular invasion decreased when HUCs were treated with MS-275. However, this was attenuated when RNASE4 and RNASE7 were silenced (Fig. 3F). These findings provide evidence that enhancing RNase 4 or RNase 7 expression through HDAC inhibition reduces UPEC susceptibility. Additionally, they suggest that HDAC class I inhibitors, like MS-275, may be candidates to pursue as RNase 4 and RNase 7 inducers given their isoenzyme specificity and reduced off-target activities compared to pan-HDACi.

MS-275 Enhances RNase 4 and RNase 7 Production In Vivo to Reduce Acute UTI Susceptibility.

To investigate whether MS-275 increases RNase 4 and RNase 7 expression in vivo, we treated humanized RNase 7 transgenic mice with MS-275. The use of RNase 7 transgenic mice allowed us to evaluate RNase 7 expression in vivo, as its expression is absent in rodents (12). Following 4 doses of intraperitoneal MS-275 (Fig. 4A), mice exhibited normal phenotypes, kidney and bladder histopathology, and immune cell profiles (SI Appendix, Figs. S6–S9). Western blot and ELISA show MS-275-treated mice had greater bladder and urinary RNase 4 and RNase 7 peptide expression compared to vehicle-treated mice, indicating that MS-275 augments RNase 4 and RNase 7 in vivo (Fig. 4 B and C).

Fig. 4.

MS-275 protects mice from acute UPEC challenge. (A) Schematic showing how female humanized RNase 7 transgenic mice were treated with intraperitoneal MS-275 (10 mg/kg) or vehicle every other day (arrows) and subjected to UTI. (B) Representative Western blots probed for RNase 4, RNase 7, acetyl-Histone H3 (lysine 27), and GAPDH from bladders of mice treated with vehicle or MS-275 (Left). Relative RNase 4, RNase 7, and acetyl-Histone H3 (lysine 27) protein production normalized to GAPDH as determined by densitometry from Western blots. Each point identifies protein expression in a different mouse bladder (n = 6 mice/group). Graphs show the mean expression and SEM. (C) Urinary RNase 4 and RNase 7 concentrations, normalized to urine creatinine (UCr), in vehicle- and MS-275-treated mice. The horizontal line indicates the median concentration in each group (n = 10 to 12 mice/group). Each point denotes RNase 4 or RNase 7 urinary concentrations in a unique mouse. (D) Urine isolated from vehicle- (circles) and MS-275-treated (squares) mice was incubated with and without an anti-RNase 4 antibody, anti-RNase 7 antibody, or an irrelevant antibody (IgG) prior to UPEC inoculation. The number of UPEC colony forming units (CFU) were determined after 90 min of incubation. The horizontal line indicates the median CFU of each group (n = 8 mice/cohort). Asterisks indicate significant P-values for the indicated pairwise comparison (Kruskal-Wallis). (E) Vehicle- and MS-275-treated female mice were transurethrally infected with UPEC (strain UTI89). Twenty-four hours post infection, urine, and bladders were collected, and UPEC colonies were enumerated. UPEC burden was combined from three independent experiments. The horizontal line indicates the geometric mean. Each point denotes UPEC burden in a unique mouse. (B, C, and E) Asterisks indicate significant P-values for the indicated pairwise comparison (Mann–Whitney U test). *P < 0.05, **P < 0.01, ****P < 0.0001.

To test whether boosting urinary RNase 4 and RNase 7 concentrations augments the urine’s antimicrobial capacity, we collected urine from RNase 7 transgenic mice treated with vehicle or MS-275 and inoculated it with UPEC. Compared to urine collected from vehicle-treated mice, UPEC survival decreased in urine isolated from MS-275-treated mice (Fig. 4D). To validate the antimicrobial contributions of RNase 4 and RNase 7, we added RNase 4 and RNase 7 neutralizing antibodies to separate urine aliquots from MS-275-treated mice. The addition of antibodies directed against RNase 4 or RNase 7 increased UPEC survival. The addition of an irrelevant antibody had no effect (Fig. 4D). These ex vivo data complement our in vitro findings and show that augmenting urinary RNase 4 or RNase 7 concentrations reduces UPEC survival.

To determine whether MS-275 administration impacts UTI susceptibility, 6- to 8-wk-old female RNase 7 transgenic mice were treated with four doses of MS-275 and transurethrally infected with UPEC (Fig. 4A). Twenty-four hours after infection, MS-275-treated mice had an 82% reduction in urinary and bladder UPEC burden compared to vehicle-treated mice (Fig. 4E). Similar results were observed in wild-type C57BL/6 female mice treated with MS-275 (SI Appendix, Fig. S10A). To determine whether MS-275 promotes UPEC clearance 7 d post infection, we utilized a UTI superinfection model and transurethrally infected female C57BL/6J mice twice with UPEC. We used this approach as mice on a C57BL/6J genetic background are resistant to chronic infection with the single bacterial inoculum used in the previous experiments (25, 26). Seven days after the second infection, we enumerated low, but comparable, UPEC titers in vehicle and MS-275-treated mice (SI Appendix, Fig. S10B). These data suggest that MS-275 treatment augments immune defenses in vivo to acutely reduce UPEC susceptibility and this effect may not be sustained at later time points.

To investigate the impact of MS-275 treatment on cellular responses to UPEC, we profiled immune cell populations in UPEC-infected bladders using flow cytometry. Results show comparable natural killer cell, neutrophil, macrophage, and T cell profiles in bladders of mice treated with four doses of vehicle- and MS-275-treated mice, suggesting that MS-275 treatment does not impact acute immune cell recruitment to infected bladders (SI Appendix, Fig. S9).

MS-275 Alters Histone Acetylation on the RNASE4 and RNASE7 Promoters.

Histones undergo posttranslational modifications that impact their DNA binding and transcriptional regulation. Histone H3 acetylation at lysine 27 (H3K27ac) is a modification that relaxes the chromatin structure and facilitates transcription (27, 28). Our results show that MS-275 increases H3K27ac (Figs. 2 B and F and 4B). To determine whether there are regions of the RNASE4 and RNASE7 promoters associated with H3K27ac, we accessed the Encyclopedia of DNA Elements (ENCODE) Project, which contains publicly available data from chromatin immunoprecipitation (ChIP)-seq analyses (29). We identified two regions for H3K27ac in the promoters for each gene (Fig. 5A). Next, we performed ChIP to determine whether HDAC inhibition increases H3K27ac at the RNASE4 and RNASE7 promoters. We pretreated HUCs with MS-275 and performed ChIP using antibodies directed against H3K27ac, an irrelevant IgG of the same isotype as a negative control, and a Histone H3 antibody as a positive control. Data were analyzed by quantitative PCR using primers targeting the two regions for H3K27ac we identified in the RNASE4 and RNASE7 promoters (Fig. 5A).

Fig. 5.

HDAC3 is required and sufficient for RNASE4 and RNASE7 suppression. (A) Schematic of the human RNASE4 and RNASE7 genes and relative locations targeted by primers used for chromatin immunoprecipitation (ChIP) assays. Primer targets are depicted by boxes labeled with one or two. The transcriptional start site is denoted by the arrow. (B) ChIP assays were performed using human bladder urothelial cells treated with vehicle (circles) or MS-275 (squares). An anti-acetyl histone H3 (lysine 27) antibody was used to immunoprecipitate chromatin-bound DNA, and qRT-PCR was performed using primers specific to the two promoter regions of RNASE4 and RNASE7 shown in (A). Graphs show the mean and SEM of precipitated DNA compared to total input as calculated in the Methods. Results are from four independent experiments. Asterisks denote significant P values for the pairwise comparisons (Student’s t test). (C and D) Human bladder urothelial cells were transiently transfected with an HDAC1, HDAC3, or a nontargeting control (NTC) siRNA pool. (C) qRT-PCR confirmed HDAC1 and HDAC3 knockdown and (D) shows relative RNASE4 and RNASE7 transcript expression. Graphs show the mean expression and SEM. Results are from five independent experiments performed in triplicate (n = 5). Asterisks denote significant P-values for the pairwise comparisons as determined by one-way ANOVA. (E) Representative Western blots probed for HDAC1, HDAC3, acetyl-Histone H3 (lysine 27), RNase 4, RNase 7, and GAPDH in human bladder urothelial cells transiently transfected with a control (empty) or HDAC3 overexpression plasmid. **P < 0.01, ***P < 0.001, and ****P < 0.0001.

When HUCs were treated with MS-275, we observed a 2.4 to 4.8-fold increase in the proportion of RNASE4 and RNASE7 promoters bound to H3K27ac (Fig. 5B). ChIP with IgG did not result in target amplification, and ChIP with a positive control Histone H3 antibody resulted in the amplification of all promoter regions assessed. These data show that MS-275 leads to enhanced H3K27ac at specific promoter regions of RNASE4 and RNASE7 and may account for how MS-275 enhances RNASE4 and RNASE7 expression.

HDAC3 Is Required and Sufficient to Suppress RNASE4 and RNASE7 Expression.

Because MS-275 shows selectivity toward HDAC1 and HDAC3 proteins, we sought to determine whether HDAC1 or HDAC3 are required to regulate RNASE4 and RNASE7 expression (30). First, we silenced HDAC1 and HDAC3 in vitro using siRNA. HUCs were transiently transfected with an HDAC1, HDAC3, or nontargeting control siRNA pool. qRT-PCR confirmed HDAC silencing (Fig. 5C). When HDAC3 was silenced, RNASE4 and RNASE7 expression increased. In contrast, silencing HDAC1 did not impact RNASE4 and RNASE7 expression, suggesting that HDAC3 is required to suppress RNASE4 and RNASE7 (Fig. 5D).

To confirm whether HDAC3 is sufficient to repress RNase 4 and RNase 7 expression, HUCs were transiently transfected with eukaryotic plasmids that drive constitutive HDAC3 expression or empty vector control. Western blot shows that HDAC3 overexpression suppressed RNase 4 and RNase 7 expression and decreased H3K27ac (Fig. 5E). These data demonstrate that HDAC3 functions to suppress RNase 4 and RNase 7 expression. In addition, they provide supporting evidence that H3K27ac is a modification that may regulate RNase 4 and RNase 7 production.

Discussion

The armamentarium of antibiotics to treat or prevent UTI is rapidly diminishing as antibiotic-resistant uropathogens become increasingly prevalent. Consequently, new strategies or nonantibiotic alternatives to treat or prevent UTI are needed. The primary objective of this project was to identify avenues that amplify RNase 4 and RNase 7 expression to create fresh approaches to mitigate UTI. To accomplish our primary objective, we employed a high-throughput screen of FDA-approved drugs and identified pan-HDACi and HDAC class I inhibitors as RNase 4 and RNase 7 inducers. Our in vitro and in vivo validation, gene silencing studies, and antibody neutralization experiments provide evidence that enhancing RNase 4 and RNase 7 expression augments UPEC killing—establishing the initial framework to develop these AMPs as antibiotic-preserving therapies.

A growing body of evidence from our research team and others suggests that deregulated AMP expression heightens UTI risk (7). Our published data show that girls with recurrent UTI have suppressed urinary RNase 4 and RNase 7 concentrations (9, 12). In addition, girls with recurrent UTI have an increased prevalence of an RNASE7 single-nucleotide polymorphism that suppresses RNase 7’s antibacterial activity and increases UPEC susceptibility (13). Thus, augmenting RNase 4 or RNase 7 production in people with recurrent UTI or populations with an increased propensity to develop more severe UTI (i.e., people with diabetes, spinal cord disorders, or pregnant women) may have a clinical impact.

Published data show that insulin, insulin-sensitizing agents like metformin, and traditional herbal extracts augment RNase 7 production (31–33). These studies support the utility of repurposing drugs to boost AMP expression and strengthen UTI defenses. To expand upon these findings, we developed a cell-based luciferase assay to identify other compounds that induce human RNase 7 as well as RNase 4 expression. By employing this high-throughput assay, we identified 94 unique FDA-approved drugs that augment RNASE4 and RNASE7 promoter activity. Of these drugs, HDACi were the most potent AMP inducers. These findings were validated in vitro using primary human cells and in vivo using humanized RNase 7 transgenic mice. To our knowledge, this is the first study to complete a high-throughput library screen to identify categories of drugs that may be repurposed to boost RNase 4 and RNase 7 expression. Results from this screen highlight the potential value of using preclinical models and high-throughput approaches to advance therapeutic discovery for UTI.

Previously published data and high-throughput compound screens have identified small molecules, hormones, and vitamins that induce the expression of other AMPs, such as cathelicidin and β-defensins, in humans and other species (34–37). In line with our findings, a body of evidence suggests HDACi, including butyrate, trichostatin A, resveratrol, and MS-275 upregulate human β-defensin 1, β-defensin 2, or cathelicidin (38–43). While this body of work supports the potential of HDACi promoting AMP production, the mechanisms responsible for HDACi-mediated induction of individual AMP genes are not well defined and it is unclear whether these mechanisms are species or cell-type specific.

Gene expression is regulated by histone acetylation, which occurs when histone acetyltransferase enzymes transfer an acetyl group to internal histone lysine residues. Histone acetylation triggers chromatin relaxation, promotes transcription factor binding to DNA regulatory elements, and activates transcription. In contrast, histone deacetylases remove acetyl groups from histone lysine residues, repressing transcription by tightening chromatin and excluding transcription factor binding. HDACi suppress the removal of acetyl groups leading to histone hyperacetylation and increased gene transcription. Our data indicate that MS-275 promotes Histone H3 acetylation at lysine 27 residing at the RNASE4 or RNASE7 promoter—suggesting that the relaxed status of local chromatin may be a mechanism responsible for MS-275 increased RNase 4 and RNase 7 expression. In support of this mechanism, when HDAC3 is overexpressed, Histone H3 acetylation at lysine 27 is reduced and RNase 4 and RNase 7 expression is suppressed.

Since the FDA approved the first HDACi to treat cutaneous T cell lymphoma (Vorinostat), HDACi use has expanded to other cancers, neurodegenerative disorders, inflammatory bowel diseases, rheumatologic diseases, asthma, and cardiovascular disease. Currently, there are over twenty HDACi in clinical trials (44). While our understanding of the utility of HDACi to eradicate microbial infections is in its infancy, HDACi have been used with varying degrees of success in vitro, in vivo, and in clinical settings to treat viral and bacterial infections. Since a pioneering publication showing valproate, a class I selective HDACi, increased murine mortality following Klebsiella pneumoniae or Candida albicans infections, subsequent publications have shown therapeutic benefit of HDACi for the treatment of human immunodeficiency virus, COVID-19, and malaria (45–47). In the gastrointestinal tract, experimental evidence shows MS-275 treatment improved cholera recovery and reduced Vibrio cholerae shedding in infected rabbits, phenylbutyrate administration reduced the severity of shigellosis, and enteropathogenic E. coli induced diarrhea in rabbits, and a randomized controlled trial showed that adjunct therapy with enteric phenylbutyrate improved outcomes in adults with shigellosis (43, 48–50). With respiratory disease, pan and class II-specific HDACi decreased Mycobacterium tuberculosis survival in human macrophages ex vivo and tuberculosis patients receiving oral adjunct therapy with phenylbutyrate ± Vitamin D had enhanced clearance of Mycobacterium tuberculosis in sputum cultures and showed more rapid clinical recovery (51, 52). Recent reviews highlight the limitations, challenges, and promise of HDACi for the treatment of inflammatory and infectious diseases (53–55).

In the context of UTI, the effects of HDAC proteins and HDACi in infection pathogenesis or prevention have not been comprehensively investigated. Data generated using the G. mellonella larvae model show that the pyelonephritis UPEC strain CFT073 increases histone acetylation and augments AMP expression (56). Urothelial HDAC6 inhibition (a Class II HDAC) prevents UPEC invasion in vitro, yet global HDAC6 deletion does not reduce UTI risk in vivo (57, 58). Here, our findings show that HDACi protect kidney epithelial cells and bladder urothelial cells from UPEC in vitro, and gene silencing studies indicate this is partly due to augmented RNase 4 or RNase 7 expression. Our findings also demonstrate that RNase 7 expressing mice and C57BL/6 mice are protected from UPEC 24 h after infection when preloaded with MS-275. Antibody neutralization experiments suggest that augmented urinary RNase 4 or RNase 7 confer UPEC protection, yet added mechanisms are likely contributing to this phenotype. These may include enhanced expression of other AMPs or activation of immune responses that have not yet been explored. Studies are ongoing to further define how HDACs and HDACi modulate immune responses in the kidney and urinary tract.

While HDACi show promise as therapeutics, lingering questions exist regarding the influence of HDAC inhibition on the quality of immune responses to pathogens. To assess whether MS-275 treatment promotes chronic UPEC clearance, we superinfected mice pretreated with MS-275 and continued treatment 7 d after infection. These experiments required the superinfection model because mice on a C57BL/6 genetic background readily resolve UTI and/or have low levels of UPEC burden 7 d after infection (26). With this approach, we did not observe therapeutic benefits with MS-275 administration. In part, these results could be a ramification of our experimental design. Although the superinfection model promotes sustained UPEC colonization, it introduces a second transurethral catheterization which may drive urothelial remodeling and impact UTI outcomes (25, 59). Alternatively, the observed findings may be a manifestation HDACi related off-target effects. Additional studies are needed in preclinical and clinical models to define the targets and safety of HDACi. Moreover, drug titration studies, pharmacodynamic profiles, and investigation optimizing the route and timing of administration for drugs like MS-275 are needed. Early evidence suggests that the benefits of HDACi on antibacterial responses in vitro and in vivo strongly depend on compound selectivity and the timing of treatment (60). Completion of these future experiments will provide a deeper understanding of the translational potential to repurpose HDACi for UTI therapy.

To conclude, our observations underscore the potential of augmenting RNase 4 and RNase 7 production for the prevention and treatment of common infections like UTI. Understanding the interplay between epigenetic regulation, histone acetylation, and AMP expression may promote the translation of these findings into strategies to induce RNase 4 and RNase 7 production and reduce antibiotic overuse.

Materials and Methods

Construction of RNASE4 and RNASE7 Promoter Luciferase Reporter Plasmids and Generation of Cell Lines Stably Expressing These Plasmids.

Human RNASE4 and RNASE7 promoters were excised from pLightSwitch_Prom plasmids (catalog numbers S722616 and S702200; Switchgear Genomics, Carlsbad, CA, USA) and inserted into the pGL4.19 vector (Promega, Madison, WI, USA). Inserts were isolated using a restriction enzyme digestion with SacI and BglII (New England Biolabs, Ipswich, MA, USA) followed by resolving on a 1% agarose gel. Bands corresponding to the size of the inserts were cut and gel purified using the QIAquick Gel Extraction Kit (Qiagen, Hilden, Germany). The destination vector was linearized using SacI and BglII and gel purified in the same way as the inserts. T4 ligase (New England Biolabs) reaction was performed to incorporate the purified inserts into the destination vector. Plasmids were bidirectionally sequenced to ensure proper insert orientation and sequence (Eurofins, Luxembourg, Germany).

HEK293 cells (American Type Culture Collection, Manassas, MA, USA) were transfected using Amaxa Nucleofector 4D in buffer SG (Lonza, Basel, Switzerland) and program CM130 with pGL4.19 vectors containing RNASE4 and RNASE7 plasmids. Stable pools were selected using the antibiotic G418 (Geneticin, Invivogen, San Diego, CA, USA). Individual clones were isolated using limiting dilution in 96 well plates and expanded. Luciferase expression was confirmed as outlined below. These reporter cells are referred to as HEK293-R4-Luc or HEK293-R7-Luc.

Luciferase Assay and Z’-Factor Calculation.

HEK293-R4-Luc and HEK293-R7-Luc cells were seeded at 10⁴ cells per well on white-walled 96 well plates (Corning, Corning, NY, USA) in phenol-free Dulbecco’s Modified Essential Medium (DMEM; Corning) with 10% fetal bovine serum (FBS). The next day, media was changed to phenol-free DMEM with 0.5% FBS. On day 3, cells were stimulated for 6 h with DMSO (negative control) or 100 nM PMA (positive control). Luciferase activity was then measured using modified luciferase assay buffer and substrate as previously described (61). Briefly, cells were lysed in 3× Firefly Assay Buffer (150 mM Tris, 75 mM sodium chloride, 3 mM magnesium chloride, 0.25% Triton X-100, 15 mM dithiothreitol, 0.5 mM coenzyme A, 0.45 mM adenosine triphosphate, and 4.2 mg/mL D-luciferin) in a volume equal to the volume of media in the wells (100 μL). Luciferase activity was measured with a microplate luminometer (Veritas Technologies, Santa Clara, CA, USA) with 1 s integration time per well. Values were recorded as relative light units (RLU, SI Appendix, Fig. S1).

Z’-factors were calculated using the following formula, where ${\sigma }_{{\mathrm{C}}^{+}}$ is the SD of the positive control, ${\sigma }_{{\mathrm{C}}^{-}}$ is the SD of the negative control, ${\mu }_{{\mathrm{C}}^{+}}$ is the mean RLU measurement of the positive controls, and ${\mu }_{{\mathrm{C}}^{-}}$ is the mean RLU measurement of the negative controls (24).

$${\mathrm{Z}}^{\text{'}}=1-\frac{3{\sigma }_{{\mathrm{C}}^{+}}+3{\sigma }_{{\mathrm{C}}^{-}}}{\left|{\mu }_{C^{+}}-{\mu }_{C^{-}}\right|}.$$

Drug Repurposing Screen.

Screening of 1,280 FDA-approved compounds was performed at University of Michigan Center for Chemical Genomics. This drug repurposing library was formed as a subset of the Bioactive Compound Library (L1700) from Selleck Chemicals. Details on the library can be found here https://www.lsi.umich.edu/science/centers-technologies/center-chemical-genomics/services/sample-libraries under Drug Repurposing Set, and the tested compounds are listed in Dataset S1.

HEK293 cells stably expressing RNASE4 or RNASE7 promoter-driven luciferase were seeded at 3,000 cells per well onto white 384 well tissue culture plates (Greiner Bio-One, Monroe, NC, USA) in 40 μL DMEM with 10% FBS using a Multidrop Combi reagent dispenser (Thermo Scientific, Waltham, MA, USA). After 72 h, culture media was replaced with 40 μL phenol red-free DMEM with 0.5% FBS using the Biomek FX 384-well programmable liquid handling instrument (Beckman Coulter Life Sciences, Indianapolis, IN, USA). After an additional 16 h, drugs were pintooled in 2 mM stock plates and added at a final concentration of 2 μM using a Thermo Scientific Multidrop Combi cassette. After 6 h of incubation with drug or vehicle at 37 °C and 5% CO₂, luciferase assays were performed by adding an equal volume of 3× Firefly Assay Buffer to the media in the wells using the Biomek FX 384-well programmable liquid handling instrument. Plates were incubated on a rotating platform at 500 rpm for 10 min, and luciferase activity was measured using a PHERAstar microplate luminometer (BMG Labtech, Cary, NC, USA). Results were reported as RLU and z-scores were calculated using the following formula:

$$\mathrm{z}=\frac{\left(\mathrm{x}-\mu \right)}{\sigma },$$

where x is the RLU value measured for an individual compound, $\mu$ is the mean RLU value measured for all samples in an assay, and $\sigma$ is the SD in RLU values measured for all samples in an assay. Z-scores greater than two were considered positive hits and were further validated (SI Appendix, Dataset S1).

Confirmation luciferase assays were performed at University of Michigan as described above. Fold changes were calculated using the following formula:

$$\mathrm{FC}=\frac{{\mu }_{\text{x}}}{{\mu }_{{\text{C}}^{-}}},$$

where FC is the fold change in promoter activity, ${\mu }_{\mathrm{x}}$ is the mean RLU measurement for an individual compound, and ${\mu }_{{\mathrm{C}}^{-}}$ is the mean RLU measurement for vehicle control.

Validation Luciferase Assays.

HEK293-R4-Luc cells or HEK293-R7-Luc cells were seeded at 1 × 10⁵ cells per well in triplicate in a clear bottom, white-walled 96-well plate in phenol red-free DMEM containing 5% FBS. After 24 h of incubation, culture media was replaced with 100 μL of fresh phenol red-free DMEM with 5% FBS containing DMSO vehicle or HDACi drugs (Selleckchem, Houston, TX, USA) at the specified concentrations. Luciferase assays were performed as outlined above. Luminescence was recorded on a Veritas Microplate Luminometer after a treatment time of 6 or 24 h (Turner Biosystems, Sunnyvale, CA, USA). Triplicate luminescence values were averaged, and treatment luminescence values were normalized to vehicle-treated cells.

In Vitro Stimulation of Human Kidney and Bladder Epithelial Cells.

Commercially available primary human bladder urothelial cells (HBLAK cells, CELLnTEC Advanced Cell Systems, Bern, Switzerland), derived from the bladder of an 80-y-old male, were cultured in a CnT-Prime Epithelial Proliferation Medium (CELLnTec Advanced Cell Systems) media (62, 63). Commercially available primary human kidney medullary epithelial cells obtained from a 3-y-old Hispanic female (Lifeline Cell Technology, Frederick, MD) were cultured in Renalife Media (Lifeline Cell Technology, as previously published (31). Once reaching 90 to 95% confluency, cells were treated with the indicated drugs. At the indicated time points, RNA was isolated from cells for qRT-PCR or cell lysates were generated for Western blot. Conditioned media were collected for ELISA or antibacterial assays.

RNA Isolation and Quantitative Real-Time PCR.

RNA was isolated using the RNeasy Plus Mini Kit (Qiagen, Hilden, Germany) following the manufacturer’s instructions. cDNA synthesis was performed using the Verso cDNA synthesis kit (Thermo Fisher Scientific, Waltham, MA, USA). qRT-PCR reactions were performed with the 7,500 Real-Time PCR System (Applied Biosystems, Carlsbad, CA, USA) as previously described (10). Reactions included cDNA corresponding to 10 ng RNA, ABsolute Blue SYBR master mix (Thermo Fisher Scientific), and the primers outlined in SI Appendix, Table S2. Relative gene expression changes were calculated using the 2^−ΔΔCT method.

SDS-Page Western Blot and ELISA Assays.

Protein extraction and SDS-PAGE Western blot were performed as previously described (9, 18). Primary antibodies used for Western blot included rabbit anti-RNase 4 (catalog number ab200717, Abcam, Cambridge, UK), rabbit anti-RNase 7 (catalog number HPA005690, Sigma-Aldrich, St. Louis, MO, USA), rabbit anti-Acetyl Histone H3 lysine 27 (catalog number 8173, Cell Signaling Technology, Danvers, MA, USA), rabbit anti-HDAC1 (catalog number 34589, Cell Signaling Technology), rabbit anti-HDAC3 (catalog number 85057, Cell Signaling Technology), and rabbit anti-GAPDH (catalog number 5174, Cell Signaling Technology). RNase 4 and RNase 7 antibody specificity were previously validated (9, 12). Commercial ELISA assays were used to measure human and mouse RNase 4 (MyBioSource, San Diego, CA, USA) and RNase 7 (Hycult Biotech, Plymouth Meeting, PA, USA) from conditioned cell culture media or mouse urine. Urinary RNase 4 and RNase 7 concentrations were normalized to urine creatinine (Oxford Biomedical, Rochester Hills, MI, USA).

In Vitro Protein Silencing and Overexpression.

AMP and HDAC silencing: siRNA targeting RNASE4 (catalog number L-021381-01-0020), RNASE7 (catalog number L-018193-01-0020), HDAC1 (catalog number L-003493-00-0005), or HDAC3 (catalog number L-003496-00-0005) or nontargeting control (catalog number D-001810-10) were purchased from Horizon Discovery (Lafayette, CO, USA) and transfected into HUCs using Dharmafect I transfection reagent as previously described (9, 12).

HDAC overexpression: Plasmid encoding Flag-tagged HDAC3 (gifts from Eric Verdin, obtained through Addgene; catalog #13820; https://www.addgene.org/13820/; RRID:Addgene_13820) or empty vector pcDNA3.1+ (Invitrogen, Waltham, MA, USA) were transfected into HUCs using Amaxa 4D-Nucleofector X Unit, P1 Primary cell Nucleofector Solution, and Nucleofector program DS-109 (Lonza, Basel, Switzerland). Western blotting was performed on total protein isolated from overexpression construct-transfected cells 72 h after transfection.

In Vitro UPEC Attachment and Invasion Assay.

UPEC attachment and invasion assays were performed on HKCs and HUCs as published (9). Cells were treated with Givinostat or MS-275 overnight and then challenged with 2 × 10⁵ CFU/well UPEC. UPEC contact with host cells was expedited by centrifugation of plates at 1,000 rpm for 3 min. After 2 h of UPEC incubation, one set of infected cells (three wells) were lysed in 0.1% Triton X-100 and plated onto LB-agar plates to enumerate the total number of UPEC per well (extracellular and intracellular). To quantify bacterial attachment, another set of infected cells (three wells) were washed and lysed in Triton X-100 and plated on LB-agar plates. The percentage of adherent bacteria was determined as the number of CFU recovered after washing divided by the total number of CFU in each well.

UPEC invasion was determined using a gentamicin protection assay (9). An additional set of infected cells (three wells) was washed and incubated for an additional 3 h in gentamicin-containing media to kill extracellular bacteria. After washing, cells were lysed and plated onto LB-agar plates. The percentage of intracellular bacteria was determined as the number of CFU recovered after cell lysis following incubation in the presence of gentamicin divided by the total number of CFU.

Chromatin Immunoprecipitation.

DNA was immunoprecipitated from HUCs treated with vehicle or 0.5 μM MS-275 for 24 h using the SimpleChIP Enzymatic Chromatin IP Kit with magnetic beads (Cell Signaling Technology) per the manufacturer’s instructions. Each immunoprecipitation reaction was carried out using 5 μg crosslinked chromatin and a primary antibody to Acetyl-Histone H3 (lysine 27) or negative (Rabbit IgG) and positive (Histone H3) controls. Quantitative PCR reactions were performed with the 7,500 Real-Time PCR System (Applied Biosystems) using 2 μL of DNA isolated (eluted in 30 μL total) from the immunoprecipitation reactions, ABsolute Blue SYBR Green master mix (Thermo Fisher Scientific), and RNASE4 and RNASE7 primers targeting histone H3 lysine 27 acetylation peaks identified using ChIP-seq data from the ENCODE Project (SI Appendix, Table S2) (29). Results are expressed as percent precipitated DNA compared to total input, and calculated as follows:

$$\begin{array}{c}\% \mathrm{Precipitated} \mathrm{DNA}=\frac{100}{2^{{(\mathrm{C}}_{\mathrm{T}} \left[\mathrm{ChIP}\right]-({\mathrm{C}}_{\mathrm{T}} \left[\mathrm{Input}\right]- {\log }_2(\mathrm{Input} \mathrm{Dilution} \mathrm{Factor}))}}.\hfill \end{array}$$

Mouse Breeding and Genotyping.

Mouse maintenance was performed in strict accordance with the Abigail Wexner Research Institute’s Institutional Animal Care and Use Committee rules and regulations. Breeding and genotyping of hemizygous RNase 7 transgenic mice was performed as published (12).

In Vivo MS-275 Dosing and Experimental UTI.

Six- to seven-week-old female C57BL/6J or RNase 7 transgenic mice were subjected to intraperitoneal injections of vehicle [5% DMSO, 30% PEG300, 65% phosphate-buffered saline (PBS)] or MS-275 (10 mg/kg bodyweight; 5% MS-275/DMSO, 30% PEG300, 65% PBS) every other day. MS-275 dosing was selected based on prior publications showing its efficacy and safety (64–66). Urine and tissue samples were collected 24 h after the final dose.

Single inoculum UTI was performed as previously described (12, 18). On the day of the fourth MS-275 injection, mice were anesthetized and transurethrally inoculated with 10⁷ CFU of UPEC. Twenty-four hours post inoculation, mice were reanesthetized and sacrificed via cervical dislocation. After sacrifice, organs were aseptically harvested and UPEC were enumerated on LB agar plates.

Murine Urine Antimicrobial Assay.

The antimicrobial activity of RNase 4 and RNase 7 in murine urine was evaluated using colony count assays as previously described with following modifications (18). Briefly, 25 μL of murine urine was inoculated with 10⁷ CFU UPEC (strain UTI89). When indicated, 0.5 μg of a polyclonal anti-RNase 4 antibody (Abcam), monoclonal anti-RNase 7 antibody (Sigma), or an IgG isotype antibody for an irrelevant peptide was added to each test isolate thirty minutes before UPEC inoculation. After 90 min of incubation at 37 °C, serial dilutions of the reaction mixtures were plated on LB agar, and the number of UPEC CFUs was determined the following day.

Methods used to generate supplemental data are provided as SI Appendix.

Statistical Analysis.

Continuous differences between groups were evaluated for a normal distribution with the D’Agostino-Pearson Omnibus or Shapiro-Wilk test, with normality defined as a P-value > 0.05. Comparisons on normally distributed data were performed by an unpaired t test or by a paired t test when appropriate; otherwise, the nonparametric Mann–Whitney U test (unpaired data) was used. Differences between groups with a P-value < 0.05 were regarded as statistically significant. Data from in vitro experiments were normally distributed and are presented as means ± SEM. Vehicle- and drug-treated cells were compared by one-way ANOVA.

Supplementary Material

Appendix 01 (PDF)

Dataset S01 (XLSX)

Data, Materials, and Software Availability

All study data are included in the article and/or SI Appendix.