ABSTRACT
Haemophilus parainfluenzae is a nutritionally fastidious, Gram-negative bacterium with an oropharyngeal/nasopharyngeal carriage niche that is associated with a range of opportunistic infections, including infectious endocarditis and otitis media (OM). These infections are often chronic/recurrent in nature and typically involve bacterial persistence within biofilm communities that are highly resistant to host clearance. This study addresses the primary hypothesis that H. parainfluenzae forms biofilm communities that are important determinants of persistence in vivo. The results from in vitro biofilm studies confirmed that H. parainfluenzae formed biofilm communities within which the polymeric matrix was mainly composed of extracellular DNA and proteins. Using a chinchilla OM infection model, we demonstrated that H. parainfluenzae formed surface-associated biofilm communities containing bacterial and host components that included neutrophil extracellular trap (NET) structures and that the bacteria mainly persisted in these biofilm communities. We also used this model to examine the possible interaction between H. parainfluenzae and its close relative Haemophilus influenzae, which is also commonly carried within the same host environments and can cause OM. The results showed that coinfection with H. influenzae promoted clearance of H. parainfluenzae from biofilm communities during OM infection. The underlying mechanisms for bacterial persistence and biofilm formation by H. parainfluenzae and knowledge about the survival defects of H. parainfluenzae during coinfection with H. influenzae are topics for future work.
INTRODUCTION
Biofilms are communities of bacteria attached to a substratum or interface, which are encased in an extracellular matrix (1). Biofilm formation is considered an adaptation of bacteria to hostile environments (1, 2). Biofilm bacteria usually have distinct characteristics compared to planktonic bacteria, including an increased resistance to antimicrobial treatments and immune system responses (3). It has been estimated that over 60% of all bacterial infections may involve biofilms (3). These biofilm-related infections include a majority of chronic and recurrent diseases and usually involve long-term bacterial persistence (3, 4). Otitis media (OM), one of the most common childhood infections (5) and the leading reason for pediatric office visits and new antibiotic prescriptions to children (6), has long been thought of as involving bacterial persistence within biofilms (1, 7, 8). Clinical evidence of bacterial biofilms includes direct observation of biofilms on the middle ear mucosa of patients with chronic OM (9) and in the chinchilla infection model of OM (10–17).
Bacteria that inhabit mucosal surfaces typically form multicellular differentiated biofilm communities that are characterized by their promotion of persistence and resistance to antibiotics (18). Haemophilus parainfluenzae is a nutritionally fastidious, Gram-negative bacterium that normally resides within the human nasopharynx and periodontal spaces, where its carriage is usually without significant symptoms (19). This species is differentiated from the closely related Haemophilus influenzae by the lack of a strict requirement for hemin and differences in its surface glycolipid populations (20–23). Notably, many of these surface factors lacking in H. parainfluenzae are those that we have previously shown to be important determinants of biofilm formation and maturation for H. influenzae (14, 24–26). Therefore, biofilm formation by this organism could be quite different from that of H. influenzae. When mucociliary clearance is compromised, H. parainfluenzae can disseminate and cause opportunistic infections that include periodontal infections, infectious endocarditis, pulmonary infections associated with chronic obstructive pulmonary disease, and OM (27–32). These diseases are often chronic/recurrent in nature and may involve bacterial persistence within biofilm communities, which are beneficial for bacterial persistence and antibiotic resistance. Recently, studies using static biofilm assays showed that H. parainfluenzae can form biofilms in vitro (33, 34); however, the structure and matrix components of the biofilms are unknown and there is no current study to test the impact of biofilm formation/maturation by H. parainfluenzae in vivo.
The chinchilla model has been widely used to study H. influenzae, Streptococcus pneumoniae, Pseudomonas aeruginosa, and Moraxella catarrhalis OM infections, establishing that these pathogens form biofilms in vivo (35). Addressing biofilms in the context of an intact immune system and the complex environmental and structural features present in a mammalian host is necessary to determine if biofilm formation is relevant to bacterial persistence in clinical infections. In this study, we first confirmed that H. parainfluenzae formed dense biofilm communities whose matrix was mainly composed of DNA and proteins. Next, using the chinchilla infection model, we demonstrated that H. parainfluenzae formed surface-associated biofilm communities containing a bacterial population, host cells, and neutrophil extracellular trap (NET) structures and that the bacteria mainly persisted in these biofilm communities. The opportunists that cause OM are normally carried within the nasopharyngeal microbiome. H. influenzae is an extremely common inhabitant of the human nasopharynx and is a leading cause of OM. In order to directly assess the polymicrobial interaction within the middle ear of the chinchilla, animals were transbullarly infected with both organisms. We found that coinfection with H. influenzae promoted clearance of H. parainfluenzae from biofilm communities. The underlying mechanisms of bacterial persistence and biofilm formation by H. parainfluenzae and the survival defects of H. parainfluenzae during polymicrobial infection with H. influenzae are topics of current study.
RESULTS
H. parainfluenzae forms biofilms in vitro.Biofilm formation by H. parainfluenzae was tested using a microtiter plate biofilm initiation assay. The results of the time course experiment, shown in Fig. 1A, demonstrated that H. parainfluenzae formed an initial biofilm at 4 h following inoculation. From 4 to 8 h, it presented early biofilm growth. At 24 h, the biofilm biomass reached the highest level and represented biofilm maturation. To further investigate the characteristics of H. parainfluenzae biofilms, we assessed bacterial communities formed in 4-well slides with LIVE-DEAD staining and imaged them with confocal laser scanning microscopy (CLSM). As shown by the results in Fig. 1B, H. parainfluenzae formed thick biofilms containing dense and multiple layers of bacteria. Viable organisms were predominant within the bacterial communities.
Characterization of biofilm formation by H. parainfluenzae in vitro. (A) Microtiter plate biofilm assay indicates biofilm formation by H. parainfluenzae. Bacteria were cultured in 96-well plates. After increasing amounts of time, biofilms were washed and quantitated by crystal violet staining. Bars represent the mean absorbance. Error bars show the standard deviations. (B) Static biofilm assay with LIVE/DEAD staining to visualize H. parainfluenzae biofilms. Bacteria were cultured in 4-well chamber slides for 24 h. Biofilms formed on chamber slides were washed and stained with LIVE/DEAD BacLight reagents and visualized by CLSM. Syto 9 staining (green) labeled viable cells, whereas propidium iodide staining (red) indicated dead cells or cells with a damaged membrane. Z-series images were used to create representative volumetric views of H. parainfluenzae biofilms. Propidium iodide staining revealed that the majority of cells within 24-h biofilms were viable.
Biofilm matrix, which holds bacterial biofilm cells together, is a complex mixture of macromolecules, including exopolysaccharides, proteins, and DNA (36). The composition of the biofilm matrix varies among bacterial species and strains (36). To define the matrix components of H. parainfluenzae biofilms, we treated biofilms with different concentrations of DNase I (Fig. 2A) or proteinase K (Fig. 2B) for 1 h to detect the detachment of biofilms. When the biofilms were treated with either DNase I or proteinase K, the biofilm density was significantly diminished in a dose-dependent manner, indicating that the biofilm matrix was likely composed of DNA and proteins. Notably, biofilms were not detached by treatment with the carbohydrate-modifying agent sodium metaperiodate (data not shown), which has been shown to detach biofilms by degrading matrix polysaccharides (37), indicating that carbohydrates are not likely to form a significant component of the biofilm matrix.
H. parainfluenzae biofilms are susceptible to detachment by DNase I and proteinase K (PK). Bacterial biofilms formed in 96-well plates were washed and incubated with different concentrations of DNase I (A) or PK (B) in enzyme buffer for 1 h at 37°C. Biofilms were quantitated by crystal violet staining as described in Materials and Methods. Bars represent the mean absorbance. Error bars show the standard deviations. Asterisks indicate significant decreases in biofilm biomass following enzymatic treatment compared to the untreated biofilm biomass. P values were determined by Student's t test. *, P < 0.05; **, P < 0.01.
During biofilm growth, bacteria produce matrix to attach bacteria together in biofilm communities. DNase I, proteinase K, or sodium periodate was added to the culture medium at the time of inoculation; the results indicated that DNase or proteinase but not periodate significantly inhibited biofilm formation by H. parainfluenzae, as determined using the microtiter plate biofilm assay (Fig. 3A). These findings suggest that the decreased biofilm biomass with enzyme treatment was mainly related to the digestion of either matrix protein or DNA but not carbohydrate. We hypothesized that matrix-degrading enzymes may affect biofilm architecture. We imaged 24-h bacterial biofilms formed in 4-well slides stained with Syto 9 to label all biofilm cells and visualized them by CLSM. Without enzyme treatment, H. parainfluenzae formed smooth biofilms without pillar and tower structures on the surface (Fig. 3B). When treated with DNase I (Fig. 3C), the biofilms with complex tower-shaped architectures on the surface were thinner than mock-treated biofilms but thicker than proteinase K-treated biofilms, which had rougher surfaces with more pillar and small tower structures on the top (Fig. 3D). No such changes in biofilm structure were observed for biofilms treated with periodate (data not shown). To confirm the CLSM observations of biofilm architecture, the COMSTAT image analysis tool was used to evaluate three variables of biofilm architecture: average biofilm thickness (Fig. 3E), total biomass (Fig. 3F), and roughness coefficient (Fig. 3G). The biofilms with DNase I treatment had significantly decreased average thickness and total biomass compared to those of mock-treated biofilms. Similarly, the biofilms with proteinase K digestion had more dramatically decreased average thickness and total biomass. Another parameter that was clearly affected by enzyme treatment was the roughness coefficient. A rough biofilm has many pillars and towers, whereas a smooth biofilm consists of more homogenous layers of cells (38). When the biofilms were treated with either DNase I or proteinase K, the biofilm roughness coefficient was significantly increased, indicating the formation of rough and heterogeneous biofilms. These results confirmed that both DNase I and proteinase K not only inhibited biofilm formation but also affected biofilm architecture. No significant changes in any structural parameter were observed following treatment with periodate, suggesting that carbohydrate is not a major component of the H. parainfluenzae biofilm matrix.
DNase I and proteinase K (PK) inhibit H. parainfluenzae biofilm formation. Bacteria were cultured with NBHI medium or NBHI medium supplemented with DNase I (1 mg ml−1) or proteinase K (1 mg ml−1) in 96-well plates, 24-well plates, or 4-well chamber slides for 24 h. (A) Biofilms formed in 96-well plates were washed and quantitated by crystal violet staining for measuring biofilm biomass. (B) Biofilms formed in 24-well plates were serially diluted and plated for viable bacterial counts. Biofilms formed in 4-well chamber slides were stained with Syto 9 alone for labeling all biofilm cells and visualized by CLSM. (C to E) Z-series images were used to create representative volumetric views of untreated (C), DNase I-treated (D), and proteinase K-treated (E) biofilms. (F to H) Z-series images were also exported to COMSTAT to obtain biofilm measurements, including average biofilm thickness (F), total biomass (G), and surface roughness coefficient (H). Error bars indicate standard deviations. Asterisks indicate a significant difference compared to the untreated biofilm as determined by Student's t test. **, P < 0.01.
H. parainfluenzae forms biofilms in the middle ear space of experimentally infected chinchillas.The chinchilla has been established as a valid infection model system for testing biofilm formation in vivo (35). The primary hypotheses for our infection studies were that H. parainfluenzae forms biofilms during experimentally induced OM and that these biofilms promote bacterial persistence in the middle ear. Chinchillas were infected via the middle ear superior bullae with either 104 or 108 CFU of bacteria. The animals were monitored for signs of OM by video otoscopy every other day. Groups were harvested on days 3, 7, and 14 postinfection. After euthanasia, the thin layer of bone comprising the superior bullar surface was excised and the interior of the bulla was observed and imaged from above. Not only effusion fluid but also thick and opaque exudate was macroscopically visible within the chinchilla middle ear chamber (Fig. 4A, right), which was similar in appearance to the biofilms formed by nontypeable H. influenzae (NTHi) and S. pneumoniae, the two most common pathogens that cause OM in humans. The results of infection studies are summarized in Table 1. The inoculum dose correlated with the frequency of signs of OM, including the presence of biofilms or effusion in the bulla or the presence of abnormal tympanic membrane, and with the results of otoscopy checks. Biofilms and effusions were observed in a majority of animals infected with 108 CFU of bacteria at all time points after infection. Only one bulla infected with 104 CFU of bacteria had visible biofilm at either day 3 or 7 after infection, and none of the animals showed effusion and visible biofilms at day 14. Abnormal tympanic membrane and/or surrounding tissue were observed in 71% of bullae in the high-dose infection group and only 10% of those in the low-dose infection group. Moreover, viable H. parainfluenzae bacteria were recovered from bullar homogenates (surface-adherent populations) in the high-dose infection group at all time points and in the low-dose infection group at days 3 and 7 (Fig. 4B). Notably, no viable bacteria were observed in middle-ear effusion fluids (planktonic populations) in the low-dose infection group at all time points (Fig. 4C). Although viable bacteria were recovered from middle-ear effusion fluids in the high-dose infection group, both the bacterial counts and the incidence of detectable bacteria were significantly lower than those from bullar homogenates (Fig. 4B and C). These results confirmed that H. parainfluenzae formed biofilms during experimentally induced OM in the chinchilla middle ear and that persistence of the bacteria was mainly from the surface-associated (biofilm) community. Taken together, these results are consistent with the biofilm hypothesis for bacterial persistence during chronic OM.
H. parainfluenzae persists in the middle ear of the chinchilla model of OM. Chinchillas were infected via the middle ear superior bullae with either 104 (circles) or 108 (triangles) CFU of bacteria. Groups were harvested on days 3, 7, and 14 postinfection. (A) Photographs show a representative middle ear chamber of a mock-infected animal (left) and an infected animal with macroscopically visible biofilm and effusion (right) at day 3 after infection. (B and C) Bacterial counts were obtained from bullar homogenates (B) and effusions (C) of infected animals. Long horizontal lines show the limit of detection. Short horizontal lines represent the geometric mean of CFU in each group. Data represent the results from three independent experiments.
Summary of chinchilla infection experiments
Bacteria and host cells are visible within chinchilla middle ear biofilms.To determine the composition of the middle ear biofilms, cryosections of biofilms obtained from the superior bullar surface of chinchillas at day 3 postinfection were stained with rabbit anti-Haemophilus polyclonal antisera coupled with Alexa 488-conjugated anti-rabbit antibody and visualized by CSLM. Fluorescent antibody staining revealed bacterial communities (Fig. 5A, left) within a matrix that was not reactive with antisera, indicating that it was not composed of bacteria or bacterium-derived materials (Fig. 5A, right). Histopathological analysis of cryosections of biofilms stained with hematoxylin and eosin (H&E) revealed that many host cells were embedded within a dense fibrous network surrounding the host cells (Fig. 5B, left) and that the majority of them had a morphology consistent with polymorphonuclear (PMN) cells (Fig. 5B, right). The presence of PMN cells associated with a fibrous mesh is similar to that in biofilms of NTHi, P. aeruginosa, and S. pneumoniae OM infections (14, 15, 17, 39).
Visualization of H. parainfluenzae and host cells within chinchilla middle ear biofilms. Chinchillas were infected via transbullar injection with 108 CFU of bacteria and euthanized at 3 days postinfection. (A) Immunofluorescence staining visualized bacterial communities within the biofilms. Cryosections of biofilms obtained from superior bullar surfaces were stained with rabbit anti-Haemophilus polyclonal antisera coupled with Alexa 488-conjugated anti-rabbit antibody and visualized by CLSM. Bacteria within the cryosections were visible in green (left; bar, 10 μm). Different interface contrast (DIC) image (right) indicates biofilm matrix. (B) Histopathological analysis revealed neutrophils within chinchilla middle ear biofilm. H&E staining of biofilm cryosections at low magnification revealed that many host cells were embedded within the biofilm structure (left; magnification, ×20) and at high magnification that the majority of them were polymorphonuclear cells (right; magnification, ×100). (C) LIVE/DEAD staining revealed viable host and bacterial cells within chinchilla middle ear biofilms. Unfixed portions of fresh chinchilla middle ear biofilms obtained from superior bullar surfaces at 3 days postinfection were immediately stained using the LIVE/DEAD BacLight kit and visualized by CLSM. White arrow indicates viable bacteria (green, left).
In order to detect the presence of viable bacteria and host cells in the biofilm materials, unfixed portions of H. parainfluenzae biofilms recovered from the chinchilla bullar surface were immediately stained with LIVE/DEAD reagents. Confocal images showed that many viable cells were stained green with Syto 9 (Fig. 5C, left) and dead cells or cells with damaged membranes were stained red with propidium iodide (Fig. 5C, right). These cells were identified both as host and bacterial cells by size and morphology. In some fields of the confocal images, fewer viable host and bacterial cells were observed (Fig. 6A, left). Notably, viable bacteria were visible within a weblike fibrous DNA matrix stained with propidium iodide (Fig. 6A, right), which was consistent with NET structures (17, 40, 41). The defining characteristics of NET structures include the presence of a high concentration of antimicrobial components, such as histone and elastase, attached to the DNA fibers. Thus, we stained cryosections of the biofilms with antibodies that recognized both histone (Fig. 6B) and elastase (Fig. 6C). These images clearly showed colocalization of DNA with histone or with elastase within a weblike fibrous structure. Thus, H. parainfluenzae formed biofilms in the middle ear of chinchillas that contained both bacterial and host components with characteristics consistent with in vivo biofilms containing NET structures in other species.
Bacteria were visible within NET structures in the chinchilla middle ear biofilms. (A) LIVE/DEAD staining revealed viable bacteria within the fibrous DNA lattice. Unfixed portions of fresh chinchilla middle ear biofilms obtained from superior bullar surfaces at 3 days postinfection were immediately stained with LIVE/DEAD BacLight reagents and visualized by CLSM. The white arrow indicates viable bacteria within a chinchilla middle ear biofilm (green, left). Dead cells or extracellular DNA lattice was stained with propidium iodide (red, middle). Bacteria within fibrous DNA lattice structures are visible in the merged image (white arrow, right). (B and C) Immunofluorescence staining visualized eukaryotic histone (B) and elastase (C) within a fibrous DNA lattice in the chinchilla middle ear biofilms. Cryosections of chinchilla middle ear biofilms were stained with mouse anti-histone or mouse anti-elastase antibodies, followed by an Alexa 488-conjugated goat anti-mouse secondary antibody (left, green). Propidium iodide was used to costain DNA (red, middle). Merged images (right, yellow) indicate colocalization between DNA lattice and either histone in panel B (right, merge) or elastase in panel C (right, merge). Bar size, 10 μm.
Coinfection with H. influenzae promoted H. parainfluenzae clearance from biofilm communities in chinchilla model of OM.Epidemiology studies show that many cases of OM are polymicrobial in nature (42). In order to examine the possible interaction between NTHi and H. parainfluenzae, coinfection studies were performed in the chinchilla infection model of OM disease. At all time points, high numbers of NTHi were recovered and NTHi was not affected by the presence or absence of H. parainfluenzae in bullar homogenates (Fig. 7A). However, H. parainfluenzae in bullar homogenates was significantly affected in the coinfection group (Fig. 7B). At days 7 and 14 postinfection, the viable counts and incidence of detectable H. parainfluenzae in bullar homogenates were significantly reduced in the coinfection group compared to the viable counts and incidence in a single infection. Notably, H. parainfluenzae in the coinfection group was cleared from all the samples at day 14. These results indicate that coinfection with NTHi promoted H. parainfluenzae clearance within the surface-associated communities (biofilms).
Polymicrobial infection promoted H. parainfluenzae clearance from biofilm communities in chinchilla model of OM. Chinchillas were infected with 103 CFU of NTHi 86-028 NP, 108 CFU of H. parainfluenzae, or a mixture of both species. Animals were harvested on days 3, 7, 14, and 21. After middle ear effusion fluids were removed, bullae were washed with PBS, removed, and homogenized for enumeration of viable NTHi (A) and H. parainfluenzae (B) cells by plating on SBHI or NBHI medium, respectively, with added vancomycin. Black circles represent CFU recovered from single-species bullar homogenates. Black triangles represent CFU recovered from polymicrobial-species bullar homogenates. Long horizontal lines show the limit of detection. Short horizontal lines represent the geometric mean of CFU in each group. Statistical significance was assessed by Mann-Whitney nonparametric analysis. *, P < 0.05 compared to the number of CFU from single-species bullar homogenates.
DISCUSSION
Biofilms are generally defined as communities of bacteria attached to a substratum or interface that are encased in an extracellular polymeric matrix material and exhibit resistance to exogenous stressors (4, 43, 44). Recently, H. parainfluenzae biofilm formation was observed in vitro using a microtiter plate assay with crystal violet staining (33, 34); however, the structure and components of H. parainfluenzae biofilms remain unknown. The results in the present study clearly demonstrated that H. parainfluenzae formed smooth biofilms containing dense and multiple layers of viable and dead bacteria without pillar and tower structures on the surface. The biofilm matrix that holds biofilm cells together varies between bacterial species and strains and is a complex mixture of macromolecules that can include exopolysaccharides, proteins, and DNA (45). Our results indicated that the H. parainfluenzae biofilm matrix is likely composed of DNA and proteins, based on the findings that both DNase I and proteinase K inhibited biofilm formation, affected biofilm architecture, and detached preformed biofilms. Notably, biofilms were not detached by treatment with the carbohydrate-modifying agent sodium metaperiodate (data not shown), which has been shown to detach biofilms depending on matrix exopolysaccharides (37, 46), indicating that carbohydrates are not likely to form a significant component of the H. parainfluenzae biofilm matrix. Biofilm bacteria usually have distinct characteristics compared to planktonic bacteria, including an increased resistance to antimicrobial treatments (4). We also observed that H. parainfluenzae biofilm cells exhibited highly increased resistance to killing by antibiotics compared to the susceptibility of planktonic cells (data not shown). Taken together, our in vitro results determined that H. parainfluenzae biofilms exhibited phenotypes similar to those of many other bacterial biofilms, including viable and dead bacteria, a protein and DNA matrix, and increased antibiotic resistance. The data in this study support the conclusion that H. parainfluenzae forms biofilms in vitro. Further studies are needed to identify determinants of biofilm formation by H. parainfluenzae.
Bacteria that inhabit mucosal surfaces typically form multicellular biofilm communities that are characterized by their promotion of persistence (47). Addressing biofilms in the context of an intact immune system and the complex environmental and structural features present in a mammalian host is necessary to determine if biofilm formation is relevant to bacterial persistence in clinical infections. However, there is no study characterizing biofilm formation by H. parainfluenzae in vivo at present. In this study, the chinchilla infection model of OM, one of the useful animal models for studying in vivo biofilms, especially as they pertain to OM, was used to investigate whether H. parainfluenzae formed biofilms in vivo. The results in the present study clearly demonstrated that H. parainfluenzae formed surface-associated bacterial communities containing viable and dead bacteria and host cells and matrix components, including NET structures, as has been observed for biofilm formation by other species. Further research will be necessary to determine the contribution of host cells and NET structures that can inhibit or possibly facilitate H. parainfluenzae biofilm formation. Additionally, viable bacteria were mainly recovered from the surface-associated (biofilm) communities. This study and others support the hypothesis that biofilm formation can promote bacterial persistence in the context of infectious disease.
Although epidemiology studies show that many cases of OM are caused by polymicrobial infection by opportunists that reside within the nasopharynx, we found that coinfection with NTHi promotes clearance of H. parainfluenzae from biofilm communities in OM infection, which seems clinically relevant, given that NTHi, but not H. parainfluenzae, is a leading cause of OM. Notably, no difference in the viability of NTHi and H. parainfluenzae in both single and polymicrobial biofilms was observed in vitro. Taken together, these data suggested that NTHi promoted clearance of H. parainfluenzae from biofilm communities in OM infection, possibly by priming enhanced host immune responses to clear H. parainfluenzae. The underlying mechanisms of the survival defects of H. parainfluenzae during coinfection with NTHi are being studied in our laboratory. Further studies are also necessary to investigate the possible interaction between H. parainfluenzae and NTHi in the nasopharynx, where both organisms share the same niche.
MATERIALS AND METHODS
Bacterial strains and culture conditions.H. parainfluenzae strain ATCC 33392 was obtained from the American Type Culture Collection. Bacteria were cultivated in brain heart infusion (BHI) medium (Difco) supplemented with 10 μg/ml NAD (Sigma) at 37°C and 5% CO2 (48). This medium is referred to below as NAD-supplemented BHI (NBHI). Nontypeable H. influenzae (NTHi) strain 86-028NP is a nasopharyngeal isolate from a child with chronic OM (49) for which an annotated complete genomic sequence is available (50). NTHi 86-028NP was cultivated in BHI medium supplemented with 10 μg/ml hemin (ICN Biochemicals) and 10 μg/ml NAD. This medium is referred to below as supplemented BHI (SBHI).
Static biofilm initiation.The in vitro biofilm formation analysis was performed with a well-described microtiter assay (26, 51). Overnight cultures of H. parainfluenzae from NBHI plates were diluted to ∼107 CFU/ml in NBHI broth, inoculated into 96-well microtiter dishes (106 CFU in 100 μl/well), and incubated at 37°C and 5% CO2. At various times thereafter, the plates were removed from the incubator, washed, and stained with 1% crystal violet (26). The remaining crystal violet in the wells was solubilized with ethanol and quantified by measurement of the optical density at 540 nm (OD540).
For the biofilm detachment assay, biofilms were cultured for 24 h in 96-well microtiter dishes as described above. Biofilms were rinsed with water and treated with 100 μl of 50 mM Tris-HCl (pH 8.0) containing 1 mg/ml proteinase K (Sigma) or 100 mM NaCl and 1 mM CaCl2 containing l mg/ml DNase I (Sigma) for 1 h at 37°C, and then total biomass was quantified by crystal violet staining as previously described (52).
Static biofilm assay.The static biofilm assay was performed essentially as described previously (25, 53). Briefly, approximately 107 bacteria were inoculated into Lab-Tek II no. 1.5 four-chamber German cover glass slides (Nunc). After incubation at 37°C and 5% CO2 for 24 h, the medium was removed and biofilm samples were washed with phosphate-buffered saline (PBS) and stained with LIVE/DEAD BacLight viability reagents (Invitrogen). Biofilms were visualized by confocal laser scanning microscopy (CLSM) using a Nikon Eclipse C1 CLSM microscope; vertical Z-series images were used to compare biofilm structure and density according to standard methods (38).
For the biofilm inhibition assay, bacteria were cultured in 96- or 24-well plates or 4-well chamber slides for 24 h as described above, except that NBHI broth was supplemented directly with 1 mg/ml proteinase K or DNase I; controls were cultured in NBHI broth only. Biofilms formed in 96-well plates were washed and quantitated by crystal violet staining for measuring biofilm biomass. Biofilms in 24-well plates were serially diluted and plated for viable bacterial counts. Biofilms in 4-well chamber slides were stained with Syto 9 alone for labeling all biofilm cells and analyzed by CLSM as described above, and the results quantitated with COMSTAT analysis as follows: the Z-series images were visualized using NIS-Elements AR 3.1 software and exported as tagged-image-format (TIF) files into the MATLAB package (The Math Works, Inc., Natick, MA). Six to nine Z-series image stacks, each representing a different field of view, were compiled and analyzed in the Image Processing Toolbox as previously described (38, 54). Measurements of total biomass and average thickness and surface roughness coefficients were chosen to characterize the biofilm structures.
Chinchilla infection studies.Bacterial persistence and biofilm formation in the middle ear were assessed essentially as described in our previous work (14, 24). Chinchillas (5/group) were purchased from Rauscher's chinchilla ranch (Larue, OH) and allowed to acclimatize to the vivarium for at least 7 days prior to infection. All animals were assessed by a veterinarian at intake and confirmed to have no overt disease symptoms at the time of infection. The animals were anesthetized with isoflurane and infected via transbullar injection with either ∼104 or ∼108 CFU of H. parainfluenzae in single-infection experiments and with ∼108 CFU of H. parainfluenzae, ∼103 CFU of NTHi 86-028NP, or a mixture of both species in polymicrobial infection experiments as indicated for each experimental group. Bacterial loads within all inocula were confirmed by plate counts. The establishment and progression of OM was assessed by otoscopic examination at 48-h intervals; the severity of inflammation was tracked by quantification of disease scores for 7 different criteria associated with OM disease. Animals exhibiting severe inflammation or signs of discomfort (head tilt, lethargy, and anorexia) received analgesia; animals with severe symptoms over >2 days were removed from the study and euthanized. Animals were euthanized at different days postinfection by cervical dislocation and exsanguination as indicated. After euthanasia, the superior bullae were aseptically opened to expose the middle ear cavity as described previously (14, 16, 17), and the presence of visible biofilm formation was assessed. If present, middle ear effusion fluids were collected. The middle ear cavity was lavaged with 1 ml of sterile PBS. Effusion and lavage fluids were combined, serially diluted, and assessed by plate count. For enumeration of tissue-associated bacteria, the middle ear superior bullae were excised and homogenized in 10 ml sterile PBS using a PowerGen 1000 homogenizer (Fisher Scientific); the bullar homogenates were plated to assess tissue-associated bacterial loads. All of the chinchilla infection protocols were approved by the Wake Forest and UAB Institutional Animal Care and Use Committees.
Viability staining.Visible biofilms were excised from the upper middle ear chambers of chinchillas immediately following euthanasia as described previously (17, 40, 55). Unfixed portions of fresh chinchilla middle ear biofilms obtained from the superior bullar surface at 3 days postinfection were immediately cut into small pieces (2 to 3 mm), incubated with l ml of PBS containing a mixture of Syto 9 and propidium iodide for 15 min, and then visualized by CLSM.
Cryosectioning, immunofluorescence, and histopathological examination.Additional portions of the biofilms obtained from superior bullar surfaces were processed for immunofluorescence and hematoxylin and eosin (H&E) staining. Samples were fixed in 2% paraformaldehyde overnight. After several rinses with PBS, the samples were put into Cryomold (Sakura Finetek USA, Torrance, CA), OCT compound (Sakura Finetek USA, Torrance, CA) was added, and the blocks were frozen at −70°C. Serial 5-μm sections were cut with Accu-Edge low-profile blades (Feather Safety Razor, Japan) at −20°C and stored at −70°C.
For immunofluorescence staining, the slides were brought to room temperature and then fixed briefly with 2% paraformaldehyde prior to antibody and DNA staining. For detection of H. influenzae in biofilms, individual cryosections of biofilms were stained with rabbit anti-Haemophilus polyclonal antisera coupled with Alexa 488-conjugated anti-rabbit antibody (Invitrogen, Oregon). For detection of histone or elastase, cryosections were stained with mouse anti-histone antibody (monoclonal antibody H11-4/MAB3422; Chemicon) or mouse anti-elastase antibody (monoclonal antibody Sc55548; Santa Cruz Biotech), followed by an Alexa 488-conjugated goat anti-mouse secondary antibody (Invitrogen, Oregon), and costained with propidium iodide for DNA as indicated in the text. Stained samples were analyzed using CLSM.
For histopathological examination, cryosections were stained with H&E, mounted with Permount (Electron Microscopy Sciences), and examined on a Nikon Eclipse microscope, in accordance with standard protocols that we have described previously (56).
Statistics.Statistical significance was determined by analysis of variance coupled with the post hoc t test or by Mann-Whitney nonparametric analysis as indicated in the figure legends. A P value of <0.05 was considered to be significant. Analyses were performed using GraphPad Prism, version 5 (GraphPad Software, San Diego, CA).
ACKNOWLEDGMENTS
We acknowledge excellent assistance by Uma Ghandi and Stephen H. Richardson for technical assistance with the chinchilla experiments. Staff of the WFUHS Animal Research Program provided invaluable assistance with infection studies.
This work was supported by grants from NIH/NIDCD (grant numbers DC007444 and DC10051) and a study contract from AstraZeneca. We have no potential conflicts of interest to disclose.
FOOTNOTES
- Received 28 December 2016.
- Returned for modification 29 January 2017.
- Accepted 28 June 2017.
- Accepted manuscript posted online 3 July 2017.
- Copyright © 2017 American Society for Microbiology.
