Urethral Catheter Biofilms Reveal Plasticity in Bacterial Composition and Metabolism and Withstand Host Immune Defenses in Hypoxic Environment

Biofilms composed of multiple microorganisms colonize the surfaces of indwelling urethral catheters that are used serially by neurogenic bladder patients and cause chronic infections. Well-adapted pathogens in this niche are Escherichia coli, Proteus, and Enterococcus spp., species that cycle through adhesion and multilayered cell growth, trigger host immune responses, are starved off nutrients, and then disperse. Viable microbial foci retained in the urinary tract recolonize catheter surfaces. The molecular adaptations of bacteria in catheter biofilms (CBs) are not well-understood, promising new insights into this pathology based on host and microbial meta-omics analyses from clinical specimens. We examined catheters from nine neurogenic bladder patients longitudinally over up to 6 months. Taxonomic analyses from 16S rRNA gene sequencing and liquid chromatography–tandem mass spectrometry (LC-MS/MS)–based proteomics revealed that 95% of all catheter and corresponding urinary pellet (UP) samples contained bacteria. CB biomasses were dominated by Enterobacteriaceae spp. and often accompanied by lactic acid and anaerobic bacteria. Systemic antibiotic drug treatments of patients resulted in either transient or lasting microbial community perturbations. Neutrophil effector proteins were abundant not only in UP but also CB samples, indicating their penetration of biofilm surfaces. In the context of one patient who advanced to a kidney infection, Proteus mirabilis proteomic data suggested a combination of factors associated with this disease complication: CB biomasses were high; the bacteria produced urease alkalinizing the pH and triggering urinary salt deposition on luminal catheter surfaces; P. mirabilis utilized energy-producing respiratory systems more than in CBs from other patients. The NADH:quinone oxidoreductase II (Nqr), a Na+ translocating enzyme not operating as a proton pump, and the nitrate reductase A (Nar) equipped the pathogen with electron transport chains promoting growth under hypoxic conditions. Both P. mirabilis and E. coli featured repertoires of transition metal ion acquisition systems in response to human host-mediated iron and zinc sequestration. We discovered a new drug target, the Nqr respiratory system, whose deactivation may compromise P. mirabilis growth in a basic pH milieu. Animal models would not allow such molecular-level insights into polymicrobial biofilm metabolism and interactions because the complexity cannot be replicated

Actinobaculum massiliense Proteome Profiled in Polymicrobial Urethral Catheter Biofilms

Actinobaculum massiliense, a Gram-positive anaerobic coccoid rod colonizing the human urinary tract, belongs to the taxonomic class of Actinobacteria. We identified A. massiliense as a cohabitant of urethral catheter biofilms (CB). The CBs also harbored more common uropathogens, such as Proteus mirabilis and Aerococcus urinae, supporting the notion that A. massiliense is adapted to a life style in polymicrobial biofilms. We isolated a clinical strain from a blood agar colony and used 16S rRNA gene sequencing and shotgun proteomics to confirm its identity as A. massiliense. We characterized this species by quantitatively comparing the bacterial proteome derived from in vitro growth with that of four clinical samples. The functional relevance of proteins with emphasis on nutrient import and the response to hostile host conditions, showing evidence of neutrophil infiltration, was analyzed. Two putative subtilisin-like proteases and a heme/oligopeptide transporter were abundant in vivo and are likely important for survival and fitness in the biofilm. Proteins facilitating uptake of xylose/glucuronate and oligopeptides, also highly expressed in vivo, may feed metabolites into mixed acid fermentation and peptidolysis pathways, respectively, to generate energy. A polyketide synthase predicted to generate a secondary metabolite that interacts with either the human host or co-colonizing microbes was also identified. The product of the PKS enzyme may contribute to A. massiliense fitness and persistence in the CBs

Proteins in AUP samples are degraded to the level of peptides consistent with activities of cathepsin G, proteinase 3 and, elastase.

<p><b>(A)</b> Peptide maps for protein S100-A9, prolifin-1, and histone H2B. This data is derived from peptidome analyses combining the fractions UP_sol1, UP_sol2, and UP_sol3 from both sample #94 and sample #134. No enzymatic cleavage sites were pre-selected in the database searches. The NE, PRTN3, and CTSG specific cleavage sites determined from peptide termini are mapped along each the respective protein sequence. Peptide termini not consistent with preferred cleavage sites of the three proteases were rare. The peptides, which are highlighted in the form of red and blue bars along the amino acid sequence to mark where they end, revealed peptide clusters around common cores. <b>(B)</b> Relative abundances of peptides associated with protein localization or functional groups comparing peptidome (PEP) and equivalent shotgun proteome (PROT) datasets. Quantification of peptides is based here on peptide-spectral counts. Protein groups denoted on the right of the graphic have color codes pertaining to the following names, functions, and localizations: NG, neutrophil granules; HIST, histones; CYT, cytosol; ACT1, actin; CSK, cytoskeleton (except actin) and keratins; RBC/COAG, red blood cells and coagulation.</p

Urinary pellet lysates and solubilization of UP aggregates incubating with deoxyribonuclease I (DNase I).

<p>(A) SDS-PAGE gels with 4–12% acrylamide gradients were stained with Coomassie Brilliant Blue G-250. The left lane contains M_r standards. The protein extracts displayed here are #36, #88, and #64 (DUP samples), #20 and #118 (AUP samples with a phenotype showing complete or moderate loss of aggregation after 2 freeze-thaw cycles), and six AUP samples where DNase I treatment was required to homogenize the pellets under moderate agitation. The proteins UMOD and MPO marked in the gel image were identified by LC-MS/MS. The bars at the bottom show proteomic identifications (P_ID) of microbial species and, if available, leukocyte counts/ml in the original urine sediments. Acronyms denote the following: <i>NC</i>, neutrophil counts in a high power field per ml urine; <i>tntc</i>, leukocytes too numerous to count; Gv, Gardnerella vaginalis; Pm, Proteus mirabilis; Ec, Escherichia coli; Sa, Staphylococcus aureus; Ef, Enterococcus faecalis; Kp, Klebsiella pneumoniae; <i>n</i>.<i>d</i>., not determined. (B) Photos of UP samples prior to and after incubation with DNase I in PBS at 37°C for 15 to 60 min.</p

Characterization of Early-Phase Neutrophil Extracellular Traps in Urinary Tract Infections

<div><p>Neutrophils have an important role in the antimicrobial defense and resolution of urinary tract infections (UTIs). Our research suggests that a mechanism known as neutrophil extracellular trap (NET) formation is a defense strategy to combat pathogens that have invaded the urinary tract. A set of human urine specimens with very high neutrophil counts had microscopic evidence of cellular aggregation and lysis. Deoxyribonuclease I (DNase) treatment resulted in disaggregation of such structures, release of DNA fragments and a proteome enriched in histones and azurophilic granule effectors whose quantitative composition was similar to that of previously described <i>in vitro</i>-formed NETs. The effector proteins were further enriched in DNA-protein complexes isolated in native PAGE gels. Immunofluorescence microscopy revealed a flattened morphology of neutrophils associated with decondensed chromatin, remnants of granules in the cell periphery, and myeloperoxidase co-localized with extracellular DNA, features consistent with early-phase NETs. Nuclear staining revealed that a considerable fraction of bacterial cells in these structures were dead. The proteomes of two pathogens, Staphylococcus aureus and Escherichia coli, were indicative of adaptive responses to early-phase NETs, specifically the release of virulence factors and arrest of ribosomal protein synthesis. Finally, we discovered patterns of proteolysis consistent with widespread cleavage of proteins by neutrophil elastase, proteinase 3 and cathepsin G and evidence of citrullination in many nuclear proteins.</p></div

Detection of early-phase NETs by IF microscopy.

<p>AUP aliquots were paraformaldehyde-fixed on glass slides on the day of specimen collection and stored at 4°C until further use. IF staining was performed with an MPO-specific polyclonal antibody followed by an anti-rabbit IgG conjugate to the dye CFl-555, and counterstaining with DAPI. Oil immersion microscopy (not confocal) was used for imaging with phase contrast and in blue and red channels. (a-d) sample #142; (e-h) sample #146; (i-l) sample #151; (m-p) sample #157. Sample #142 shows evidence of lobulated nuclei and no evidence of extracellular chromatin release (a), intact granular structures and well-defined cell perimeters (b), MPO staining in accordance with intact granules (c), and no co-localized MPO/chromatin staining (d). Sample #146 has intact neutrophils, but also some cells where nuclei fill the entire cell space (e) and granules are diminished in the perimeter of cells according to staining for MPO (g). Co-localization of nuclei and MPO is visible in the cell perimeter suggesting the emerging loss of nuclear membranes (h). Sample #151 shows less regularly shaped nuclei with fainter staining in their perimeters suggesting nuclear membrane disintegration (i), and patchy granular staining as described above (k); MPO and nuclear staining with DAPI overlap (l). Sample #157 shows areas of flattened and disintegrating cells (m, n) and streaks of extracellular DNA (m) that co-localizes with MPO staining (o, p). The closed white arrows point to cells with intact nuclei and well-distributed cytoplasmic granules. Open white arrows point to cells filled with chromatin and a patchy staining pattern for granules (MPO). Open yellow arrows point to disintegrated cells releasing chromatin from nuclei that co-localizes with MPO.</p

Proteolytic degradation in extracts of AUP samples.

<p><b>(A)</b> Western Blots were performed with polyclonal antibodies specific for NE, MPO, LTF, and histone H4A. The lane numbers match fraction numbers (UP_sol1, UP_sol3, and UP_sol5) derived from samples #33, #94, and #112. The M_r standard consists of ten proteins denoted with kDa values. Green arrowheads point to M_r values observed for full-length proteins, including the heavy and light chains of MPO. Full-length NE (29 kDa) and a H4A fragment (8–10 kDa) were detected only in fractions of sample #94. LTF and MPO were represented by full length protein bands in UP_sol3 fractions. The S. aureus protein A (SpA) was detected in a M_r range corresponding to its post-translationally modified, cell wall-immobilized forms for sample #112 (red arrowheads). <b>(B)</b> Cleavage sites identified in the peptide sequences of MPO and H4A that were in agreement with the preferred P1 site specificities of the proteases NE and PRTN3. This data is deduced from peptide termini identified via LC-MS/MS from five AUP samples. The N- and C-termini of peptide segments shaded in green include experimentally generated trypsin-specific cleavage sites and PRTN3/NE-specific sites apparently formed as a consequence of the <i>in vivo</i> inflammatory process. Red arrowheads denote sequence positions resulting from protein maturation of precursors. Other arrowheads denote PRTN3 and NE cleavage sites (C-terminal to A, V, L, I, S, T, C, M); if colored black, the site was identified in three or more of the five examined AUP datasets.</p

Sequential extraction of DNA and proteins from AUP samples.

<p>(A) Samples #122 and #134 were analyzed in 0.5% agarose gels and stained with ethidium bromide. DNA standards (St) are denoted in kilobase pairs (kbp). Lane numbers 1–5 match fraction numbers UP_sol1, UP_sol2, etc. (1 μl extract each). In the gel image for #134, lane 2’ pertains to a repeated incubation step with PBS and DTT, and lane 3’ pertains to a shorter 10 min incubation step with DNase I. UP_sol3 (lane 3) represents incubation for 75 min. AUP sample #122 resisted disintegration of the pellet prior to the addition of DNase I; large size DNA was even retained even in fraction UP_sol5. The aggregate in sample #134 released DNA in a decreasing size range with each incubation step. DNase I can cleave nucleic acids to nucleosome monomers (~ 0.2 kbp). (B) Protein extracts of samples #112 and #122 visualized in SDS-PAGE gels. Lane numbers 1 to 5 match fraction numbers UP_sol1, UP_sol2, etc. Ten μl extract were used in each lane. Low M_r proteins were abundant suggesting protein degradation in AUP samples. The positions of LTF, α-defensin 1, and MPO (all identified by LC-MS/MS) are marked in the gel image.</p

Protein profiles of UP_sol fractions show molecular evidence of NET formation in UTI cases.

<p>Proteomic quantification for UP_sol3 fractions shows consistent similarities with profiles observed for <i>in vitro</i> generated NETs. Twenty-one highly abundant proteins were included in the graphic. Each colored segment in a stacked bar represents the relative amount of a protein in the total proteome of a fraction (including the UP_sol1 and combined UP_sol4/5 fractions; terms UP-s1, UP-s3, and UP-s4/5 are used on the x-axis). Each protein segment has a color corresponding to the color code displayed on the very right in the order of occurrence in the stacked bars. Protein names are UniProt short names. The bar displayed on the right (term ‘exp NETs’) represents the relative quantity of 13 proteins from a publication on <i>in vitro</i> generated NETs [<a href="http://www.plospathogens.org/article/info:doi/10.1371/journal.ppat.1006151#ppat.1006151.ref014" target="_blank">14</a>]. Black arrows on top of stacked bars denote samples in which the retention of insoluble DNA in UP fractions prior to the enzymatic digestion was high. Red arrows denote samples with partial release of DNA fragments prior to DNase incubation, which was in agreement with the gradual release of histones in the UP_sol1 fraction. Green arrows denote samples with high cytokeratin (KRT1) and/or UMOD contents. The y-axis value of 1 represents 100% of the proteome using the proteomic quantification tool MaxQuant (all identified protein quantities are listed in <a href="http://www.plospathogens.org/article/info:doi/10.1371/journal.ppat.1006151#ppat.1006151.s003" target="_blank">S3 Data</a>).</p

Microbial cell viability staining for infectious agents in AUP samples.

<p>Images are from oil immersion microscopy with the green (fluorescein) channel for sample #122, and green and red (rhodamine) channels for samples #146, #151 and #157. Image #122: AUP sample aliquot was incubated with SYTOX-green (5 μM in TBS) in the dark for 15 min, fixed on a glass slide at low heat (40°C), washed with water, and air-dried. C. albicans yeast forms are clearly visible. Red arrows point to dead cells (SYTOX-green stained), white arrows point to intact cells (unstained). Images #146 to #157: Sample aliquots were incubated with the live/dead differential staining kit (5 μM SYTO9 and 55 μM propidium iodide in TBS) in the dark for 15 min followed by centrifugation at 800 x g for 3 min, re-suspension in TBS, a 2^nd centrifugation step, and fixation with 4% paraformaldehyde for 15 min. #146: rod-shaped E. coli cells propidium iodide-stained (red arrow) are dead. #151: filamentous K. pneumoniae rods stained with SYTO9 are living cells (white arrow). Neutrophils (bright yellow stain) are surrounded by bacterial cells suggesting a failure of phagocytosis. #157: S. aureus cocci are trapped in NET-like structures with red and green dots indicating death and survival (red and white arrows, respectively). The #157 insert shows a cluster of dead bacterial cells.</p