Hard-to-heal wounds have become a growing healthcare problem, affecting up to 2% of the adult population (2.4–4.5 million people) in the US alone.1,2 The most prevalent types of hard-to-heal wound are vascular ulcers, diabetic foot ulcers (DFUs) and pressure ulcers (PUs), which share some features including prolonged inflammation, unresponsiveness of cutaneous cells to reparative stimuli and persistent infection due to formation of bacterial biofilms.1 A recent systematic review revealed that 78% of hard-to-heal wounds harboured biofilms.3,4 Once biofilms are well established, they cause prolonged inflammation, antibiotic tolerance and wound chronicity, eventually resulting in higher morbidity and medical costs.5,6,7 Biofilm management is an indispensable part of treatment.
Physical biofilm removal (for example, debridement, rigorous cleansing) is considered ideal to eliminate wound bioburden, giving the opportunity of a therapeutic window to expose bacteria living deep in biofilms.7,8 Unfortunately, currently available algorithms for identifying biofilm on wounds are somewhat indirect and subjective, hindering clinicians from delivering effective biofilm management.7,9,10 Moreover, during the early stage of biofilm development, referred to as critical colonisation, direct inspection is insufficient to identify biofilm presence.
To overcome this challenge, significant efforts have been expended to establish methods for the early identification of critical colonisation. Infrared thermography has been used for identifying latent inflammation. Bacterial gene expression analysis of wound exudate has been used to reveal virulent expression, as well as bedside evaluation of bacterial bioburden on wound surface by using a bacterial counting device.11,12,13 However, these indirect assessment methods cannot locate the biofilm on the wound surface. Based on the literature, current direct biofilm identification methods are considered clinically impractical due to the complexity of laboratory techniques (including the sample handing and the possibility of false negative results) and the time required to complete the analysis, as well as the invasiveness of tissue sampling.7 Thus, clinically applicable direct biofilm identification is required.
Recently, our laboratory has developed a technique called ‘wound blotting’ to capture molecules on the wound by employing the direct attachment of a nitrocellulose membrane to the wound surface.14 The positively charged nitrocellulose membrane attracts protein complexes which are generally negatively charged.15 The proteins of interest collected on the membrane can then be identified through staining methods. The previous study showed that wound blotting combined with ruthenium red staining successfully identified biofilms and predicted slough formation in PUs, showing predictive validity.16 Ruthenium red has been widely used to visualise the microscopic structure of acidic polysaccharides, a major component of bacterial biofilms.17,18 Furthermore, this method has good known-group validity to detect an artificially grown biofilm on the surface of an ex vivo pig skin model.16 Therefore, this simple method may be a high-potential, point-of-care biofilm evaluation tool that could be applied in clinical settings. However, this study also found wounds with negative signal results for biofilm which still showed an increase of necrotic tissue proportion.16 There are two possibilities for such a finding: first, wound blotting may have overlooked biofilm components on the wound; and second, factors other than bacterial biofilms may have led to slough formation. Therefore, it is essential to confirm the concurrent validity of wound blotting for detecting biofilms. Additionally, biofilm identification based on the ruthenium red staining process requires 120 minutes before visualisation. The process itself uses a reagent that has an unpleasant odour and is toxic to health and the environment. These problems hinder its clinical applicability as a point-of-care detection system.
Alcian blue is considered to have similar polysaccharide staining features to ruthenium red. It is a non-toxic staining reagent, odourless, and with a shorter visualisation processing time compared with ruthenium red. Because of these properties, alcian blue is a potential candidate as a ruthenium red substitute for our wound blotting method; however, no data are currently available to support this idea.
This study aimed to confirm the concurrent validity of wound blotting for biofilm visualisation and to confirm the usability of alcian blue as a substitute for ruthenium red. For these main aims, we employed native polyacrylamide gel electrophoresis (PAGE) as a reference method to detect and quantify biofilm components extracted from total bacterial mass or tissue samples.19 First, the ability of native PAGE in detecting biofilm from bacteria by in vitro study was confirmed. Second, evaluation of the ability of wound blotting to also detect and quantify biofilm growing on a wound surface and within wound tissue was conducted. For this purpose, rats were used to model different levels of biofilm distribution to compare the results of biofilm identification between microscopic observations and wound blotting. Third, the correlation of wound blotting with native PAGE in quantifying biofilm from PU necrotic tissue samples was investigated.
Methods
Extraction of colony biofilm extracellular matrices for in vitro study
Wild-type Pseudomonas aeruginosa PAO1, grown on sterile Luria-Bertani (LB) 10cm agar plates for 48 hours at 37°C, was used in an in vitro study. A 48-hour culture was required to yield a denser polysaccharide concentration than a 24-hour culture. This allowed the observation of stacked bands on the native PAGE wells which were not observed in the 24-hour culture (data not shown). To prepare the bacterial extract, the 48-hour-grown colony biofilm of PAO1 was scraped, homogenised by mixing and weighed. To extract the colony biofilm containing extracellular matrix (ECM) proteins, polysaccharides and extracellular DNA, the whole PAO1 colony mass was then diluted with 1000μl of sterile 1.5M NaCl containing a protease inhibitor (PI) cocktail (Nacalai Tesque, Japan) and then centrifuged at 5000×g for 10 minutes at room temperature.19 The supernatant was used for further analysis. Additional PAO1 grown over 24 hours on two sterile LB agar plates was used for biofilm serial extraction. The whole colony bacterial mass was scraped, weighed and diluted with 1500μl sterile phosphate-buffered saline (PBS). A 500μl aliquot of total biofilm suspension was spared for total mass dot-blot, while a 500μl aliquot of the dilution was serially extracted by homogenisation with 500μl of sterile 1.5M NaCl with PI, centrifuged at 5000×g for 10 minutes and the supernatant aspirated and stored for dot-blot analysis. The extraction process was repeated three times.
Biofilm infected cutaneous wound in animal model
Sprague Dawley male rats (seven weeks old, 210–230g, Japan SLC, Inc., Japan) were acclimated to standard housing and fed ad libitum for one week at the Animal Facility at the University of Tokyo. All rats were housed in individual cages under constant temperature and humidity with a 12-hour light–dark cycle.
Protocols of wound creation and biofilm infection model
For the creation of cutaneous wounds, the rats were anaesthetised with 2% isoflurane gas. The dorsal skin of each rat was shaved and sterilised with 70% ethanol. On each rat dorsum (left and right side), two dorsal wounds (2cm apart) were created using a 12mm diameter punch biopsy (Acu Punch, Acuderm, US) and dressed with non-adhesive hydrocellular foam dressing (Smith+Nephew, UK). Wounds were cleansed daily by irrigating with 5ml of sterile 0.9% NaCl for 5–10 seconds on each wound, gently wiped with gauze (Benri Gauze G, Suzuran Sanitary Goods Co Ltd., China) and dressed with non-adhesive hydrocellular foam dressing until post-wounding day (PWD) 4 to allow the growth of granulation tissue. At PWD 5, a bacterial inoculation was performed.
For the in vivo study, PAO1 was grown by streaking a tip of PAO1 glycerol stock in 10,000μl sterile LB broth and incubation for nine hours with constant shaking at 37°C, and centrifuged at 1790×g for 10 minutes. The LB broth was then aspirated, leaving a PAO1 pellet at the bottom of the tube. The pellet was washed with 1000μl sterile PBS three times. After pellet washing, sterile PBS was added to the tube to adjust to OD600=1.5.
The rats were divided into four groups. For the first group, three rats were used as the non-inoculation group (i.e., PBS injection). In this group, each wound was injected with 100μl sterile PBS using a 29G needle (Terumo Co., Japan) at a 15° angle into the granulation tissue. For the second group, three rats were used for the surface biofilm model, in which a 2×2cm gauze was placed on each wound surface, then 400μl of PAO1 suspension (OD600=1.5) was dripped onto the gauze. For the third group, which included two rats, each wound was injected with 100μl of PAO1 solution (OD600=1.5), using the same procedure as the non-inoculation group. For the fourth group, which used two rats, each wound received two injections (total 200μl PAO1 suspension injected) using the same procedure as the third group. After bacterial inoculation treatment, each wound was covered with a 4×4cm transparent film dressing (Tegaderm, 3M, US). On PWD 8, each wound was irrigated with 5000μl sterile 0.9% NaCl (normal saline) for five seconds before wound blotting. Then, a 2×2cm nitrocellulose membrane (Bio-Rad, US) was prewetted with sterile normal saline and blotted for 10 seconds on each wound surface. Wound blotting membranes obtained from the right-side wound of each rat were assigned for ruthenium red (Sigma-Aldrich, US) staining, while those from the left-side wounds were assigned for alcian blue (Saraya Co., Ltd., Japan) staining. All rats were then euthanised by high concentration isoflurane inhalation for wound tissue harvesting.
Biofilm matrices extraction from wound samples
For the in vivo study, a full-thickness wound section from all rats was divided into two equal parts: one part for histological analysis and the other part for biofilm extraction. In the clinical sample study, debrided necrotic tissue was collected from weekly interdisciplinary PU rounds at a university hospital from March 2015 to February 2017, and stored in 3000μl normal saline at 4°C. A total of 60 patients, with a total of 104 wounds, were recruited. From that, 33 wounds from 17 patients underwent wound debridement and samples were stored. Tissue used for analysis was selected based on its consistency (dispersed or hard tissue samples were excluded), yielding 17 tissue samples from 10 patients being included in the analysis. For biofilm extraction, wound samples were minced to be as small as possible and weighed. Samples were then added to a 500μl extraction solution (1.5M sterile NaCl with PI), homogenised, and then centrifuged at 5000×g for 10 minutes at room temperature.19
Native PAGE for biofilm identification
For the native PAGE analysis, 20µl of extracted biofilm solution was added to 20µl 2% bromophenol-blue sample buffer, then 10µl of buffered-sample was applied to each well of 4–20% pre-casting gel (Mini-PROTEAN TGX, Bio-Rad, US). After running electrophoresis with tris-glycine buffer at 200V for 30 minutes, gels were fixed with 7.5% acetic acid/20% methanol solution for 30 minutes, washed with distilled water for 15 minutes, stained with EZBlue Gel Staining Reagent (Sigma-Aldrich, US) for one hour, destained overnight with distilled water, and then scanned for analysis.
To confirm the specificity of the native PAGE analysis in identifying biofilm components, a final concentration of 100µg/ml Proteinase K (ProK) (Wako Pure Chemical Industries, Ltd., Japan) and 10,000 times diluted (1×10-3U/µl) DNase (Nippon Genetech Co., Ltd., Japan), either individually or in combination, were used to digest protein and DNA components of in vitro PAO1 biofilms, respectively, leaving polysaccharides stacked at the bottom of gel wells. For a control, distilled water was added into the PAO1 biofilm extract with the same volume as the enzyme solutions. Samples were then incubated at 37°C for two hours. After digestion, 20µl of each of the PAO1-digested biofilm extracts were spared for dot-blot staining.
Ruthenium red and alcian blue staining for wound blotting and dot-blotting
Ruthenium red staining was performed as follows: each nitrocellulose membrane (wound blotted or aliquot of supernatant dot-blotted membrane) was prewetted with distilled water, stained with 5mg/ml ruthenium red solution for two minutes, washed with distilled water, destained three times with 10% acetic acid/40% methanol solution for 30 minutes, and then scanned for analysis.20
Alcian blue staining was performed as follows: each nitrocellulose membrane was soaked in the first cation detergent solution (Saraya Co., Ltd., Japan) for 30 seconds with constant shaking, then stained with alcian blue solution for 60 seconds with constant shaking, rinsed with a second cation detergent solution for 30 seconds with shaking, and finally scanned for analysis.20 The biofilm's visualised exopolysaccharides on the nitrocellulose membrane were interpreted as positive or negative through direct visual judgement by a researcher blinded to the results of histological analyses from the in vivo study.
Anti-Pseudomonas staining
Paraffin sections of each of the rat wound samples (5μm thick) were used for immunohistochemistry. Endogenous peroxidase was inactivated with 0.3% H2O2/methanol for 30 minutes. Pseudomonas aeruginosa localisation was detected with biotin-conjugated anti- Pseudomonas antibody (Thermo Fisher Scientific, US), diluted 1:300, for 60 minutes, and the avidin–biotin complex (ABC) method (Vectastain ABC kit, Vector Laboratories, US), visualised with 3,3‘-diaminobenzidine tetrahydrochloride (DAB) (Wako Pure Chemical Industries, Ltd., Japan), and then counterstained with haematoxylin (Muto Pure Chemicals Co., Ltd., Japan).
Exopolysaccharide staining for histological samples
Biofilm exopolysaccharides were visualised by fluorescence staining. After the first dehydration, samples were incubated with 1:200 (5×10-3mg/ml) fluorescein isothiocyanate (FITC) conjugated wheat germ agglutinin (WGA) (Sigma-Aldrich, US) diluted in 1% bovine serum albumin/PBS. Exopolysaccharides on the tissue sections were visualised using a fluorescence microscope (Keyence Corporation, Japan) and their presence was judged by two researchers blinded to the wound blotting results.
Image processing
Blotted membrane signals were adjusted for colour balance by using Adobe Photoshop CS6 (Adobe Systems Incorporated, US). Blotting and native PAGE signal quantification were measured using ImageJ software (National Institutes of Health, US). After adjusting the wound blotting images into greyscale, the visualised exopolysaccharides on the wound blotting were quantified by measuring the total grey value of the blotted area using free-hand selection on ImageJ, then normalised with tissue mass.
Ethical considerations
All protocols from the in vivo study were approved by the Animal Experiment Committee, the University of Tokyo (P14-087). Permission for the analysis of the clinical samples obtained from pressure ulcer rounds was granted by the Ethics Committee of the Graduate School of Medicine, the University of Tokyo (#3757-(5)). Patient data were collected from medical records.
Statistical analyses
All statistical analyses were performed using SPSS v.23 (IBM, US). Biofilm presence confirmed by histological analysis or wound blotting was compared to calculate sensitivity and specificity. To assess the correlation between native PAGE band intensity and signal intensity from dot-blotting or wound blotting, Pearson's correlation coefficients were calculated.
Results
In vitro study
To determine whether biofilm exopolysaccharides could be stained by ruthenium red or alcian blue from wound blotting, an in vitro study was performed. The result showed that 48-hour-grown PAO1 developed biofilms that could not be digested by ProK and/or DNase, as observed by remaining stacked bands at the bottom of all gel wells (Fig 1a). Enzyme-digested biofilm extracts could be stained by both alcian blue and ruthenium red (Fig 1b, 1c). A serial biofilm extraction highlighted a gradual reduction of biofilm signal on dot-blotted membranes stained by ruthenium red and alcian blue (Fig 2a, 2b) which also corresponds with native PAGE analysis (Fig 2c).
In vivo study
In the series of wound models, representative histological results for biofilm-positive and biofilm-negative cases are shown in Fig 3. In the Pseudomonas aeruginosa-positive case with infected wound appearance (Fig 3a), localisation of inflammatory cells was notably found within the wound tissue (Fig 3b, white arrowheads). On the other hand, the Pseudomonas aeruginosa-negative case with slough on the wound surface (Fig 3e) shows inflammatory cell localisation only on the superficial wound bed (Fig 3f, white arrowheads). Biofilm presence was also confirmed with fluorescence imaging, indicated by the green arrowheads (Fig 3d, 3h). The Pseudomonas aeruginosa-positive case was confirmed by the observation of green fluorescence on top of the wound bed (Fig 3d, green arrowheads) which was absent in the Pseudomonas aeruginosa-negative case (Fig 3h).
Based on histological analysis, nine biofilm-positive results were found on and/or under the wound surface from the right-side wounds (allocated for ruthenium red staining) (Table 1) and nine from the left side (allocated for alcian blue staining) (Table 2). The sensitivity for ruthenium red was 88.9%, and 100% for alcian blue. Fig 4 describes how the biofilm's presence was determined, either positive or negative, by a researcher blinded to the group allocation. For quantitative assessment of biofilm identification, dot-blot signal intensity from tissue extract measured by native PAGE analysis showed correlation coefficients of r=0.73 (p<0.05) for ruthenium red, and r=0.70 (p<0.05) for alcian blue of both dot-blot stains (Fig 5a, 5c). Correlation coefficients between signal intensity for wound blotting were r=0.67 (p<0.05) and r=0.67 (p<0.05) for ruthenium red and alcian blue, respectively (Fig 5b, 5d).
Table 1. Biofilm indentification by histological analysis and wound blotting with ruthenium red staining
| Biofilm histological analysis (number of wounds) | ||||
|---|---|---|---|---|
| Positive | Negative | Total | ||
| Wound blotting | Positive | 8 | 0 | 8 |
| Negative | 1 | 1 | 2 | |
| Total | 9 | 1 | 10 | |
Table 2. Biofilm indentification by histological analysis and wound blotting with alcian blue staining
| Biofilm histological analysis (number of wounds) | ||||
|---|---|---|---|---|
| Positive | Negative | Total | ||
| Wound blotting | Positive | 9 | 1 | 10 |
| Negative | 0 | 0 | 0 | |
| Total | 9 | 1 | 10 | |
Clinical sample study
In order to understand the concurrent validity of wound blotting in the clinical samples, wound blotting was tested using debrided tissue samples obtained from patients with PUs. Patient and wound characteristics are summarised in Table 3 and Table 4, respectively. Representative cases with strong, medium and weak band intensity are shown in Fig 6a, 6b and 6c. Scatter plots of signal intensity of biofilms from debrided tissue extracts and wound blotting showed r=0.75 (p<0.05, Fig 6d) and r=0.77 (p<0.05, Fig 6e) for ruthenium red and alcian blue, respectively.
Table 3. Patient demographic data (n=9)
| Result | |
|---|---|
| Gender, female, n (%) | 5 (55.6) |
| Age, years, mean±SD | 61.8±16.0 |
| Disease (multiple comorbidities), n (%) | |
| Neurology | 4 (44.4) |
| Urology | 4 (44.4) |
| Respiratory | 3 (33.3) |
| Oncology | 3 (33.3) |
SD—standard deviation
Table 4. Wound characteristics data (n=17*)
| Result | |
|---|---|
| Wound size, cm2, median (IQR) | 13.5 (3.6–21.6) |
| DESIGN-R score, median (IQR) | 24 (18–26) |
| Wound location (n=10), n (%) | |
| Trochanter | 2 (20) |
| Coccyx | 2 (20) |
| Leg | 2 (20) |
| Others | 4 (40) |
Discussion
This study showed concurrent validity with reference to native PAGE analysis for identifying and quantifying biofilm components through in vitro, in vivo and clinical investigations, as well as histological analysis for visualising biofilms in the in vivo study. Furthermore, it was found that biofilms could be visualised by wound blotting stained using alcian blue with a simplified procedure, compared with ruthenium red.
To confirm whether native PAGE analysis can be used as a reference method for biofilm identification, enzyme treatments by ProK or DNase were used, and it was found that they could not digest the extracted biofilm ECMs, which indicates the stacked bands in the gel (Fig 1a) represent components other than protein or DNA, such as polysaccharides. The main interest in the present study was the common polysaccharides found in biofilms. Therefore, the specific enzymes for each of those polysaccharides (alginate, Pel and Psl) were not necessary.21,22 Instead, confirmed concentrations of ProK and DNase were used, and all enzyme-treated extracts were still detected by native PAGE, leaving polysaccharides, the other known major components of biofilms.
Biofilm identification can be performed using various methods, most commonly microscopy. Although considered as the biofilm identification gold standard, microscopic analysis may overlook biofilm presence due to its heterogenous distribution within wound tissue, and high dependence on the sampling acquisition and processing protocols.23,24 Native PAGE, however, is considered to be suitable as a biofilm detection reference in this study because it detects total biofilm components through sample extraction, allowing a vast biofilm structure from the whole wound tissue to be concentrated within the extraction solution. By using native PAGE as a reference, signals were successfully detected regardless of enzyme treatments, which suggests the potential ability to detect biofilm components by ruthenium red and alcian blue staining systems.
In order to understand whether wound blotting can also detect biofilms in different distributions (on and under the wound surface), a rat model was used. In vivo studies are indispensable, especially in understanding the holistic biofilm–host interaction contributing to a biofilm distribution which may not be observed in an ex vivo model. The animal models showed extensive PAO1 biofilm coverage on and under the wound surface, as visualised by anti-Pseudomonas and fluorescence imaging corresponding with an inflammatory response confirmed by H&E staining (Fig 3). Nonetheless, the in vivo study was not able to create a biofilm-negative model as was supposed to be observed in the PBS injection group, which might be due to the normal microfloral contamination of rat skin. Moreover, using Pseudomonas aeruginosa mutant species that do not produce biofilms as a negative control could possibly highlight the significance of the biofilm signal produced by PAO1. However, such a strain of Pseudomonas aeruginosa is not available because the production of biofilm components is regulated by a highly complex system.25 Alternatively, a biofilm serial extraction was performed to highlight biofilm signals stained by ruthenium red and alcian blue through gradual biofilm reduction. This finding supports the suggestion that biofilm signals produced by ruthenium red and alcian blue staining systems correspond to biofilm presence as it was gradually eliminated through extraction.
Using these models with positive or negative biofilms on the wound surface indicated that alcian blue had 100% sensitivity and ruthenium red had 88.9% sensitivity. Unfortunately, the study was not able to estimate its specificity due to the lack of biofilm-negative wounds. To overcome this problem, the correlation of signal intensity between native PAGE and dot-blot, or wound blotting, from ruthenium red and alcian blue, was assessed and a good correlation was indicated.
The in vivo study was an acute infection wound model, which meant that biofilm development on the wound was expected to be less in quantity and quality compared with clinically hard-to-heal wounds. Furthermore, Mendes et al. reported that DFUs were 83.7% polymicrobial.26 These multispecies biofilms were found to pose more potent pathogenic impact, which then raises the question whether it is also constructed by different matrix compositions.27 Nevertheless, the wound blotting method could still detect biofilms grown on it. Therefore, wound blotting could possibly be used for biofilm detection in clinical practice. Thus, the investigation was extended to include wound blotting validation using debrided tissue samples from patients with pressure ulcers. In clinical settings, infected hard-to-heal wounds have diverse bacterial colonisations, which may produce different biofilm components.
In this clinical sample study, it was also difficult to find wound tissues with complete negative biofilm presence to be used as controls for comparison, due to the small sample size. Similar to the in vivo study, correlation analysis of signal intensities between native PAGE and wound blotting was used to confirm the concurrent validity. Scatter plots of debrided tissue blotting intensity had good correlations for both ruthenium red and alcian blue. Additionally, although wound blotting can detect biofilms, at least on the wound surface, it can also capture the whole picture of biofilm bioburden within the tissue. This ability might be due to the abundant invasion by biofilms within necrotic tissue. As shown in the animal study, the presence of biofilm matrices is also seen below the wound surface (Fig 3e). Nevertheless, wound blotting with alcian blue staining is potentially applicable to detect biofilms in clinical settings.
As for the clinical implications, it is worthwhile to emphasise the benefits offered by this biofilm detection method. Biofilm can reach its full maturity within 2–4 days, depending on the species and growth conditions, and it is extremely resistant to biocides (for example, antibiotics and antimicrobials).28 It also can recover from mechanical disruptions such as debridement to form a mature structure within 24 hours, allowing less than a 24-hour therapeutic window for antimicrobials to be effective.28 Such rapid timing and repetitive evaluation of biofilms cannot be performed through the ‘gold standard’, which is microscopic observation on the wound tissue samples. This evaluation can result in keeping the wound critically colonised. Biofilm visualisation by wound blotting only takes two minutes with a simple noninvasive procedure. These merits allow clinicians to evaluate the effectiveness of wound cleansing procedures (for example, rigorous washing and sharp debridement) if the biofilm eradication has been made optimally at each wound care visit/treatment.
The wound blotting method also has some potential limitations. First, it may cause some levels of discomfort during the membrane attachment on the wound surface, especially if used on acute wounds. This can be minimised by attaching the membrane carefully and gently. Second, wound blotting cannot distinguish the types of pathogens living within the biofilms. However, this does not limit the potential clinical applicability of wound blotting because pathogen identification should be a part of the routine culture procedure in the clinical settings before determining appropriate antibiotic treatment. Last but not least, there is a possibility of inter-observer variability due to the handling procedure used by the evaluating clinicians, as also might be the case for other laboratory observations.
Further studies should be carried out to investigate the clinical effectiveness of biofilm-based wound management with the use of wound blotting in an intervention study.
Limitations
In this study, we only employed a single species of pathogen commonly found on hard-to-heal wounds, Pseudomonas aeruginosa. The potential biofilm identification ability of wound blotting for combinations of species of bacteria and fungi remains unclear.
Secondly, the clinical samples in this study were nonviable necrotic tissue that does not contain any healthy wound sections. This is because acquiring samples from viable tissue is clinically contraindicated. Therefore, the potential for the presence of biofilm on other healthy wound sections/areas is unknown.
Conclusion
This study provides evidence of the concurrent validity of wound blotting in identifying and quantifying biofilm on wounds compared with native PAGE or histological analysis by in vitro, in vivo and clinical investigations. The sensitivity of wound blotting was high for both ruthenium red and alcian blue. Wound blotting staining by alcian blue had a good correlation with native PAGE, equal to ruthenium red. Considering alcian blue is safe and requires only two minutes before biofilms can be visualised, for clinical application, wound blotting with alcian blue staining would be a promising measure to guide clinicians to offer biofilm-based wound management.
Reflective questions
- How could biofilm identification by wound blotting improve wound healing in the future?
- How can wound blotting also detect the presence of fungal biofilms on wounds?
- What are the intra- and inter-rater reliabilities of the wound blotting method in detecting wound biofilms in clinical settings?