Pressure ulcers (PUs) are localized injuries to the skin and underlying tissue that primarily result from unrelieved pressure. PUs are a complex and significant health problem (Chen, Ostrowski, Obenaus, & Zhang, 2009; Salcido, Popescu, & Ahn, 2007) because they are painful and difficult to treat. PUs are associated with increased risk of mortality among long-term bedfast patients, as well as increased frequency and duration of hospitalization (Health Quality Ontario, 2009). PUs differentially affect the quality of life of patients who are prone to immobility, including elderly adults and persons who are neurologically impaired, chronically hospitalized, or have spinal cord injury (Kruger, Pires, Ngann, Sterling, & Rubayi, 2013). PUs burden society with the expense of care.
Many nursing approaches to prevention and treatment of PUs have been used, including pressure redistribution devices, complying with turning schedules (McInnes, Jammali-Blasi, Cullum, Bell-Syer, & Dumville, 2013), dressings (Clark et al., 2014; Santamaria et al., 2013), and negative pressure wound therapies (Health Quality Ontario, 2006); clinical effectiveness of these interventions is inconclusive, and PU occurrence continues to be high. For example, in an extensive U.S. survey, prevalence ranged from 12.3% in healthcare facilities to 33% in patients with spinal cord injury in the community (VanGilder, Amlung, Harrison, & Meyer, 2009). Point prevalence at three Irish teaching hospitals was 18.5% (Gallagher et al., 2008). The estimate of PU prevalence in Canadian healthcare settings was reported at 26% (Woodbury & Houghton, 2004). In our view, continued high rates of occurrence and challenges in treatment of PUs reflect limited knowledge related to the cellular and molecular mechanisms of early-stage PU; thus, pathophysiological research on PUs focusing on molecular targets for promising interventions becomes a top priority.
Tissue ischemic injury has traditionally been regarded as the main factor in the development of PU (Daniel, Priest, & Wheatley, 1981; Kosiak, 1959). However, ischemia–reperfusion (I/R) injury has been considered as the major contributing determinant in the formation of PU in recent studies (Loerakker, Manders, et al., 2011; Peirce, Skalak, & Rodeheaver, 2000; Stadler, Zhang, Oskoui, Whittaker, & Lanzafame, 2004; Tsuji, Ichioka, Sekiya, & Nakatsuka, 2005). I/R injury has been defined as cellular injury resulting from the reperfusion of blood to previously ischemic and oxygen-deprived tissue. Reperfusion of blood to previously ischemia and oxygen-deprived tissue can initiate extensive cellular damage through pathways that lead to leukocyte activation and oxidative stress (Grace, 1994; Reid, Sull, Mogford, Roy, & Mustoe, 2004; Taylor & James, 2005). Upon reperfusion, leukocytes become activated and interact with the endothelium through rolling, adherence, and transmigration into the interstitial compartment; this interaction invokes an inflammatory cascade, causing cellular edema and tissue damage (Mustoe, O’Shaughnessy, & Kloeters, 2006). Meanwhile, the overproduction of reactive oxygen species triggers a process termed oxidative stress, which overwhelms the endogenous antioxidant defense system, causing oxidative injury to surrounding tissues (Toledo-Pereyra, Toledo, Walsh, & Lopez-Neblina, 2004). Myeloperoxidase (MPO), a kind of oxidative enzyme found in zurophilic granules of human neutrophils (Klebanoff, 2005), is released during inflammation and could be a key maker related to inflammatory response and oxidative injury (Welbourn et al., 1991; Zouaoui Boudjeltia et al., 2004). Another indicator of oxidative stress is malondialdehyde (MDA), which is a stable end product of the peroxidation of membrane lipids by reactive oxygen species (Avci, Akarslan, Erten, & Coskun-Cevher, 2014). Some studies showed that the presence of excessive concentrations of activated matrix metalloproteinase-9 (MMP-9) might impede the healing process in chronic wounds by destroying the growth factors and extracellular proteins essential for PU healing. MMP-9 is essential for normal tissue remodeling and is involved in inflammatory conditions (Gumieiro et al., 2013). However, no study has explored the role of MMP-9 in the formation of early-stage PU.
In addition to the inflammatory and oxidative processes, the initial metabolic event during tissue ischemia is disruption of energy metabolism by depletion of adenosine triphosphate (ATP) for active ion pumps in cells and depletion of endogenous antioxidants because of anoxia (Mustoe et al., 2006), which may result in mitochondrial dysfunction and initiate the translocation of Bax—a proapoptotic member of Bcl-2 family—from the cytosol to the outer mitochondrial membrane, thus activating apoptotic pathways. The Bcl-2 family, which is constituted by a group of proteins, has been implicated as the major regulators of the programmed cell death machinery (Hou, Cymbalyuk, Hsu, Xu, & Hsu, 2003). Bcl-2 antagonizes apoptosis, whereas Bax mediates apoptotic cell death (Hou et al., 2003; Lindqvist, Heinlein, Huang, & Vaux, 2014). Previous studies (Jiang et al., 2012; Siu et al., 2009) have tested apoptotic-related factors of PU and showed that apoptosis was involved in different processes of pressure ulceration (either early stage or healing stage). However, the specific apoptotic pathways and how they work in the pathogenesis of early-stage PU have not yet been clear until now.
Recent research has focused on hypoxia-inducible factor-1(HIF-1) in that it plays a cytoprotective role under hypoxic conditions through regulating the expression of oxygen-regulated genes, which can mediate adaption to hypoxia (Semenza, 2004; Wenger, 2002). Functional HIF-1 is a heterodimer composed of HIF-1α and HIF-1β subunits. Many studies—primarily on the heart (Maloyan et al., 2005), wound healing (Hong et al., 2014), and deep tissue injury (Sari et al., 2015)—have shown that HIF-1 plays a dual role in mediating the prosurvival and proapoptotic responses, depending on the severity of hypoxia (Chen et al., 2009). In mild hypoxia, HIF-1 initiates the expression of adaptive genes, whereas in severe hypoxia, it induces proapoptotic gene expression (Chen et al., 2009). These findings led us to hypothesize that HIF-1α functions, either as a proapoptotic factor or cell protector, during the hypoxic condition of early-stage PU formation.
Because of the ethical limitations, we definitely cannot carry out fundamental biological studies to explore these hypotheses on humans; thus, an animal model mimicking clinical early-stage PU becomes the optimal choice (Renn & Dorsey, 2011). Although a few models (Peirce et al., 2000; Salcido et al., 2007) have incorporated I/R injury into wound formation, these investigations just adopted high pressures and/or a limited number of I/R cycles to create acute PUs, which undermined the clinical relevance.
The purpose of this study was to (a) develop a simple rat model of early-stage PU induced by cyclic I/R according to clinical setting, then, based on this model, (b) characterize the HIF-1α expression profile and its potential role during the early stage of PU, and (c) elaborate the apoptotic pathways and the interaction of relevant factors, such as HIF-1α gene expression, oxidative stress, inflammatory response, and energy depletion, in order to explore the cellular and molecular mechanisms of early-stage PU.
Materials and Methods
The research involved two experiments. The purpose of Experiment 1 was to establish an animal model of early-stage PU (Stage I PU). Then, using the established animal model, the purpose of Experiment 2 was to study the mechanisms for pathophysiological changes induced by I/R injury in the development of early-stage PU.
Male adult Sprague–Dawley rats (~300–350 g) were randomly caged within the cages. They were seriously maintained and cared for under approved guidelines, with a protocol approved by Institutional Animal Care and Use Committee of Sichuan University. Before starting the experiments, all rats were housed in pathogen-free conditions at 20°C, in a cycle of 12 hours of light and 12 hours of dark, and were fed rat chow and water ad libitum. After acclimatization with the housing environment, they were subjected to the pressure protocols.
The compression apparatus was designed simply according to the fundamental principle of the formation of PUs. As can be seen in Figure 1, it consisted of five main parts, including two trestles, two saucers, two pressure pillars, an electronic scale, and a foam carpet. The saucer was connected with the pressure pillar, which can deliver pressure to the limbs of rats—with a 1.0-cm diameter of pressure area. On the top of the saucer, weights can be added to produce a given pressure. The two limbs of a rat can be simultaneously pressurized with this device, and the net magnitude of pressure can be obtained through the final reading on the electronic scale minus the weights of foam carpet and rat.
Each rat was anesthetized by intraperitoneal injection with 10% chloraldurate at 300 mg/kg body weight; then, the hair of the rats’ limbs was depilated with shavers. At appropriate intervals, supplemental doses of anesthetic were administered to maintain unconsciousness. At the end of each pressure session, each animal received an intraperitoneal injection of physiological 2.5% dextrose and 0.45% saline to maintain adequate hydration. At the end point of each group, after collecting skin and muscle samples, rats were euthanized by injecting an overdose of chloraldurate.
I/R Injury Procedure
In Experiment 1, a total of 48 rats were randomly divided into six equal groups using a table of random numbers; cage was ignored during the randomization. The specific pressure protocol is illustrated in Table 1. One cycle consisted of 2 hours of ischemia and 0.5 hours of reperfusion, which meant that each cycle included 2 hours being pressurized and 0.5 hours of pressure relief. This is clinically relevant because it is recommended that patients at risk for PU be manually repositioned at certain times (e.g., two hourly turning; McInnes, Jammali-Blasi, Bell-Syer, Dumville, & Cullum, 2011).
In Experiment 2, 36 rats were randomly divided into three groups (Table 1). Group G was the control group with no I/R cycles. Group H and Group I were assigned to receive the same magnitude of pressure (based on findings from Experiment 1; 170 mm Hg), but different cycles of I/R: Group H completed 3 I/R cycles, receiving a total of 6 hours of ischemia and 1.5 hours of reperfusion; Group I completed 5 I/R cycles, receiving a total of 10 hours of ischemia and 2.5 hours of reperfusion. At the completion of each group, samples were collected for later analyses; then, animals were euthanized.
Macrographs Before and After Compression in Experiment 1
Macrographs of the treatment tissue before and after compression were taken using a digital camera (Samsung NX100, Seoul, South Korea). Appearance in the change of the skin among all groups can be observed, such as skin color, skin shape, and skin integrity. As nonblanchable erythema is defined by the National Pressure Ulcer Advisory Panel as the typical symptom of clinically Stage I PU (Black et al., 2007), the occurrence of nonblanchable erythema was taken as the indicator for successfully establishing the model.
Full-thickness biopsies (including the muscle layers) were obtained from the compressed region for histological analysis in both Experiments 1 and 2. All samples were fixed in 10% buffered formalin, embedded in paraffin, sectioned perpendicular to the skin surface at 5 μm thickness, and stained with hematoxylin and eosin for light microscope observation. The histological analysis was performed to determine the presence of any degenerative histological characteristics.
MPO Analysis by Enzyme-Linked Immunosorbent Assay
Muscle tissue collected from Experiments 1 and 2, which were previously snap frozen in liquid nitrogen and stored at −80°C, were triturated into homogenate using micro tissue grinders (IKA, Staufen, Germany), after adding 0.01 M precooling PBS, pH 7.4, 0.01 M PBS at the weight-to-volume ratio of 9:1. After centrifugation (20 minutes at 3000 rpm), the supernatant was collected to test the quantity of MPO using the rat MPO enzyme-linked immunosorbent assay (ELISA) kit, according to the manufacturer’s standard instructions (R&D Systems, Minneapolis, MN). Briefly, the procedure included the dilution of standard variety, adding sample, incubation at 37°C for 30 minutes, washing, adding enzyme, incubation again, coloration, and finally stop of reaction. The color change is measured spectrophotometrically at a wavelength of 450 ± 10 nm. The concentration of MPO in the samples is then determined by comparing the optical density of the samples to the standard curve.
MDA and MMP-9 Analyses by ELISA
In Experiment 2, muscle tissue previously stored at −80°C was first prepared for homogenate using the procedure described above. After collection of the supernatant, the content of MDA and MMP-9 were examined by kits, respectively (Rat MDA ELISA Kit and Rat MMP-9 ELISA Kit, R&D Systems). Actual density of MDA and MMP-9 were calculated using the same method as that of MPO.
ATPase Activity Test by Spectrometry
In Experiment 2, Na+/K+ ATPase activity, Ca2+/Mg2+ ATPase activity, and total ATPase activity were measured with the spectrophotometer colorimetric method. Compressed muscle tissue from experimental groups and the same part tissue in the control group were first ground into homogenate (on ice) with precooled normal saline at a weight-to-volume ratio of 9:1—after centrifuged (10 minutes, at 1500 rpm)—collected the supernate, then the homogenate was diluted to 2%, adding normal saline at a volume ratio of 1:4. After completion of all reactants according to instructions for the ultramicro ATPase assay kit (Nanjing Jiancheng Bioengineering Institute, Nanjing, China), the absorbance of all test tubes was monitored at a wavelength of 636 nm by ultraviolet spectrophotometer (U-0080D, Hitachi, Tokyo, Japan). Finally, three kinds of ATPase activity (Na+/K+ ATPase activity, Ca2+/Mg2+ ATPase activity, and total ATPase activity) were calculated respectively via the following formula:
where ATPase activity was measured in U/mg prot and OD is the optical density.
Terminal dUTP Nick-End Labeling Analysis
In Experiment 2, apoptotic muscle nuclei were assessed by terminal dUTP nick-end labeling (TUNEL). A fluorometric TUNEL detection kit was used according to the manufacturer’s instructions (11684817910, In Situ Cell Death Detection Kit, POD, Roche, Mannheim, Germany). After completing the standard TUNEL procedure, TUNEL-stained nuclei were examined under a fluorescence microscope equipped with a digital camera (FV1000, Olympus, Tokyo, Japan). The numbers of TUNEL-positive nuclei were counted, and data were expressed as TUNEL index, which was expressed as the percentage of TUNEL-positive nuclei. For each muscle, it was calculated from five random, nonoverlapping fields at magnification of ×400 (fluorescence microscope) or ×100 (light microscope).
Bcl-2 and Bax Proteins Analyses by Immunohistochemistry
The expression of apoptotic factors, Bcl-2 and Bax proteins, was examined by immunohistochemical staining using a streptavidin-peroxidase (SP) detection method (Xi et al., 2011) in the compressed and control uncompressed tissues in Experiment 2. Sections were incubated with rabbit anti-rat Bcl-2 (1:200; ab7973, Abcam, Cambridge, MA) and Bax (1:200; ab7977, Abcam, Cambridge, MA) antibodies at 4°C overnight. Tissue was incubated with goat anti-rabbit IgG (1:200; PV-6001, Beijing Zhongshan Goldenbridge Biotechnology, Beijing, China) at 37°C for 30 minutes. Cells were colored with DAB. Nuclei were counterstained with hematoxylin, followed by gradient ethanol dehydration, xylene transparency, and neutral gum mounting. Finally, positive immunohistochemical results, usually expressed the appearance of brown–yellow, were observed under a light microscope (×100) from five random, nonoverlapping fields of each group and then quantitatively analyzed to obtain an integral optical density (IOD) value using Image-Pro Plus 6.0 software (Media Cybernetics, Rockville, MD): The higher IOD value, the more Bcl-2 and Bax proteins.
Real-Time Polymerase Chain Reaction Gene Expression Analysis for HIF-1αα
For Experiment 2, the expression of HIF-1α was measured on total RNA extracts using an SYBR green-based real-time polymerase chain reaction according to the manufacturer’s protocol (Applied Biosystems, Foster City, CA) in a total volume of 20 μl. The procedure was performed in Real Master Mix/20× SYBR solution, primer, cDNA and RNase, and DNase-free water on an ABI 7900 real-time PCR machine (Applied Biosystems). Primer sequences were designed using the Primer Express 2.0 software (Applied Biosystems) as follows: HIF-1α, 5′-TGCTTGGTGCTGATTTGTGAA-3′,5′-TATCGAGGCTGTG TCGACTGAG-3′; β-actin,5′-TCTGTGTGGATTGGTGGCTCTA-3′, 5′-CTGCTTGCTGATCCACATCTG-3′. The 20-μl reaction mixture consisted of 9.0-μl Real Master Mix/20× SYBR solution, 0.5 μl of each primer (10 μM), 2.0 μl of sample cDNA, and 8.0 μl double distilled water. The mixture denatured at 95°C for 60 seconds and was subjected to 40 cycles of 95°C for 10 seconds and 60°C for 30 seconds, then 80 cycles of 95°C for 1 minute, 55°C for 1 minute, 55°C for 10 seconds and 4°C. Relative quantification was performed by using the comparative 2−ΔΔCt method (Livak & Schmittgen, 2001) and Light Cycler Data Analysis software (Roche). In brief, Ct value indicates the cycle number at which a significant increase in ΔRn (i.e., fluorescence) is first detected. A sample will be considered positive at the cycle in which the change in the fluorescence exceeds an arbitrary threshold value. Comparative Ct calculations for genes of interest were expressed relative to the β-actin signal generated from the same cDNA sample. The average Ct for the gene of interest was subtracted from the corresponding averaged β-actin Ct value for that sample to give a ΔCt value. The ΔΔCt values were achieved by subtracting the control ΔCt value from the compressed ΔCt value. Data were expressed as the fold differences over controls normalized to the housekeeping gene.
Means and standard deviations were reported. One-way ANOVA was used to test differences of MPO, MDA, MMP-9, ATPase, HIF-1α gene, and apoptotic factors between control and experimental groups. Nominal Type 1 error rate was set at .05 for all statistical tests. Pearson correlations were used to index linear relationships between these factors.
We first established an animal model of Stage I PU and then studied mechanisms by which I/R injury produces Stage I PU. Through the analyses, we found that apoptotic factors and MMP-9 worked together in the formation of early-stage PU. As I/R cycles increased, apoptotic muscle nuclei were elevated (p < .01) and were negatively correlated with the Bcl-2/Bax ratio but was positively correlated with MMP-9 (p < .01). Another interesting result was that HIF-1α first decreased in the 3 I/R group but increased significantly in the 5 I/R group (p < .05). And it may function in association with apoptosis during the hypoxic condition of early-stage PU development, which can be shown by the negative correlation with Bcl-2 (p < .05). Detailed results follow.
By comparing the appearance change between groups before and after compression, Control Group A had normally reactive intact skin (Figure 2A), whereas Group D (170 mm Hg) showed nonblanchable erythema (Figure 2B), which was similar to a clinical Stage I PU. As the pressure increased to 270 mm Hg, hyperemia was aggravated (Figure 2C).
Our histological analysis showed that dermis tissue and muscle tissue, following the compression procedure, generally showed characteristics of inflammation and degeneration (Figure 3). Consistent with a previous report, the morphological damage in muscle tissue was heavier than that in cutaneous tissue in the formation of early-stage PU (Kwan, Tam, Lo, Leung, & Lau, 2007). The structure of dermis tissue in Control Group A was clear, without any inflammation (Figure 3A), and muscle fiber lines were in alignment with clear striation (Figure 3B). The compressed tissue from Group B to Group F showed degenerative characteristics. These included rounding-shaped myofibers, atrophied or fractured myofibers, edema, infiltration of neutrophils, accumulated numbers of nuclei in the interstitial space, fibroplasia, centralized nuclei in the myofibers, and even autolysis (Figure 3C–P).
The MPO quantity tested by ELISA in Experiment 1 was increased in groups with more pressure. In Control Group A, MPO was little expressed, whereas in Group D in which pressure was 170 mm Hg, the MPO was significantly higher (p <.01; Table 2).
Conclusion: Experiment 1
Taken together, an animal model of Stage I PU was successfully established based on findings from analysis of macrographs, tissue, and biochemistry. A pressure at 170 mm Hg was shown as the optimum value for this model.
MPO, MDA, MMP-9 Analyses by ELISA
The examination of MPO, MDA, and MMP-9 by ELISA showed the same pattern of differences. They were remarkably higher in the muscle tissue following 5 I/R compressions than in the 3 I/R group and control group. Quantities of MPO and MDA in the 5 I/R group were slightly higher than those in the 3 I/R group (p < .01) and the control group (p < .01). The quantity of MMP-9 was slightly higher in the 3 I/R group compared to that in the control group, whereas it was significantly higher in the 5 I/R group (p < .05; Table 3).
Total ATPase activity, Na+/K+ ATPase activity, and Ca2+/Mg2+ ATPase activity in muscle tissue were tested by spectrometry among groups. The activity of the three kinds of ATPases were significantly decreased when I/R cycles were increased from 0 to 3 and then to 5. The total ATPase activity was 1.629 ± 0.004 U/mg prot in the control group, whereas it remarkably decreased to 1.365 ± 0.004 U/mg prot in the 3 I/R group and finally dropped to 1.055 ± 0.049 U/mg prot in the 5 I/R group. Na+/K+ ATPase and Ca2+/Mg2+ ATPase activity showed the same variation tendency (Figure 4).
Apoptotic muscle nuclei were measured among groups using TUNEL staining (Figure 5A–F) and were expressed as TUNEL index (Table 3). We took fluorescent photos and usual light microscope photos to clearly identify apoptotic muscle-associated nuclei. It indicated that the apoptotic index in the compressed muscle tissue was significantly elevated, relative to the control muscle tissue (p < .01; Table 3).
The Protein of Bax and Bcl-2 Expression
The protein of Bax and Bcl-2 was examined by immunohistochemical staining and was analyzed with IOD. In Control Group G (Figure 6A), there was nearly no expression of Bax protein, whereas in 3 I/R Group H (Figure 6B), it increased compared with the control group (p < .01). In 5 I/R Group I (Figure 6C), it expressed more than that of the former two groups (p < .01).
The expression of Bcl-2 protein showed a different tendency. In 3 I/R Group H (Figure 6E), it was significantly elevated compared to the control group (Figure 6D, p < .01), whereas in 5 I/R Group I (Figure 6F), the expression of Bcl-2 was significantly reduced compared to that of the former two groups (p < .01).
Gene Expression of HIF-1α
The expression of HIF-1α mRNA was estimated by real-time polymerase chain reaction analysis. It was found that HIF-1α mRNA expression in 3 I/R Group G decreased a little, compared to Control Group G, but the difference was not significant (p > .05). As I/R cycles increased from three to five, it increased significantly (p < .05; Table 3). The melt curve (see Figure, Supplemental Digital Content 1, http://links.lww.com/NRES/A161) illustrated that HIF-1α mRNA and β-actin were the only amplicons of the amplification products.
MDA was positively correlated with MPO (r = .90, p < .001), MMP-9 (r = .83, p < .001), and the apoptosis index (AI; r = .92, p < .001) but was negatively correlated with total ATPase (r = −.91, p < .001). AI was positively correlated with MMP-9 (r = .75, p < .001) but was negatively correlated with the Bcl-2/Bax ratio (r = −.64, p < .05). HIF-1α had negative correlation with Bcl-2 (r = −.46, p = .005). No significant linear relationship between HIF-1α and MDA was detected (r = .25, p = .20); scatterplots are shown (see Figure, Supplemental Digital Content 2, http://links.lww.com/NRES/A162).
PUs comprise a serious and challenging health problem, for there exists wide variability and complexity in the pathophysiology. Understanding the possible mechanisms and signal events that regulate the initiation and development of early-stage PU may provide valuable information for potential targets of intervention. In this study, we showed that HIF-1α was activated—which cooperated with the underlying apoptotic pathways, during the development of early-stage PU on a feasible animal model. In Experiment 1, we showed that three cycles of 2 hours of compression pressure at 170 mm Hg followed by 30 minutes of reperfusion produced local tissue changes (micrographs, histology, neutrophil release of MPO). In Experiment 2, we then used this level of pressure (170 mm Hg) and increased the number of compression/reperfusion cycles to show increasing tissue levels of biomarkers indicative of oxidative stress (MPO, MDA), inflammation (MMP-9), ischemia (ATPase activity, HIF-1α), and apoptosis (TUNEL, Bax, Bcl-2).
As for the magnitude of pressure and duration in this study, we referenced previous published models (Daniel et al., 1981; Nola & Vistnes, 1980; Peirce et al., 2000; Salcido et al., 2007), as well as the clinical-relevant parameters. Since Salcido et al. (2007) had stated that the mean blood pressure of experimental rats was 120 mm Hg, we chose the pressure range from 100 to 270 mm Hg and included enough time (three cycles of I/R) and finally found that 170 mm Hg was the most appropriate pressure for producing Stage I PU in our animal model; because of the occurrence of typical nonblanchable redness and inflammatory response, such as inflammatory cell infiltration, accumulated nuclei in the interstitial space of muscle tissue, as well as the increased MPO, which are attributable to intravascular hemoconcentration, leukocyte blocking, and platelet aggregation (the so-called “no-reflow” microscopic phenomenon).
PU can be initiated in cutaneous or deep tissues (Salcido et al., 2007). Because muscle tissue has lower tolerance for mechanical compression compared to other soft tissues, previous research has implicated it as the site of initial insult in the development of PUs (Bouten, Oomens, Baaijens, & Bader, 2003; Daniel et al., 1981; Gefen, 2008). Our histological analysis results confirmed that muscle tissue changed ahead of skin in the pathogenesis of early-stage PU as well.
A few publications have indicated that apoptosis is involved in the development of PUs (Gawlitta et al., 2007; Siu et al., 2009; Wang, Bouten, Lee, & Bader, 2005) or in the process of wound healing (Jiang et al., 2012), but how apoptosis functions in the formation of early-stage PUs is still unclear. A merit of the present findings is that we comprehensively tested inflammatory responses, oxidative stress reaction, apoptosis, and cell energy depletion, which may interact in the process of I/R injury (Blaisdell, 2002; Loerakker, Oomens, et al., 2011). Our data showed decreases of three kinds of ATPase activity, increases of MPO, MDA, and AI, as well as involvement of Bcl-2 family proteins. This suggests that the initial hypoxia induces dysregulation of ATPase and excess neutrophil adhesiveness and perpetuates the generation of oxygen free radicals via a neutrophil oxidative burst, which finally results in mitochondrial dysfunction and initiates mitochondria-mediated apoptotic pathways through the regulation of Bcl-2 family proteins. The ratio of Bcl-2/Bax decreased while AI became elevated when I/R cycles were increased, and they were negatively correlated. The expression of Bax protein increased after 3 I/R cycles, which we interpreted to mean that the antiapoptotic effect of Bcl-2 was exceeded, after which the affected tissue progressed to apoptosis. This also implied that the ratio of Bcl-2/Bax could be a predictive indicator for apoptosis. Furthermore, our data showed that MMP-9, a kind of proteinase degrading extracellular matrix, increased during the process of I/R injury and had positive correlations with MDA and AI. These results suggested that MMP-9, up-regulated by oxidative stress reaction, may be associated with the tissue injury through destroying growth factors and extracellular matrix proteins, initiating “anoikis” apoptosis (i.e., apoptosis induced by inadequate or inappropriate cell–matrix interactions; Frisch & Screaton, 2001), and finally damaging the integrity of skin and muscle tissue, as well as vascular wall, which could be partly the reason for the formation of PU. Although many previous studies of MMP-9 focused on chronic wound healing (Ladwig et al., 2002) and cancer (Shen et al., 2014), none has explored the specific role of MMP-9 in the development of early-stage PU. Additional research is warranted to completely dissect the specific mitochondria-mediated apoptotic signaling pathways, the potential role of MMP-9, and oxidative stress factors that may affect the sequential apoptotic events in the etiology of PU.
We also found that HIF-1α gene expression was elevated after five cycles of I/R. This is opposite to findings from studies on brain injury, which showed that HIF-1α expression was immediately up-regulated in the initiation of hypoxic injury (Umschweif et al., 2014; Wu et al., 2014). We inferred two possible reasons for this phenomenon. First, the process of I/R injury in PU, which involves incomplete and indirect ischemia followed by incomplete and chronic reperfusion, is much different from the I/R injuries in other organs (liver, brain, heart; Aras et al., 2013; Rao et al., 2014; Weymann et al., 2014). Second, cerebral cells are more sensitive and susceptible to ischemia and hypoxia than muscle tissue. Therefore, the up-regulation of HIF-1α expression in muscle tissue requires a certain period of incomplete I/R cycles—at least five cycles in our model. However, some studies reported a cytoprotective role forHIF-1α in chronic wounds (Sisco, Liu, Kryger, & Mustoe, 2007) and muscle tissue (Ameln et al., 2005) when suffering hypoxia. This seemed different from our result in that HIF-1α expression was negatively correlated with Bcl-2 protein. We interpreted this to mean that the decreased expression of HIF-1α in three cycles of I/R may work to promote muscle cell apoptosis, whereas the increasing Bcl-2 proteins simultaneously tried to resist apoptosis. The exact apoptotic pathway induced by HIF-1α and the upstream mechanism for the activation of HIF-1α in the development of PU need to be further elucidated.
The current study is the first in vivo study to show that the HIF-1α gene is activated and associated with apoptosis, in response to chronic I/R injury during the formation of early PU, which provides valuable information for molecular targets of prevention and therapy. However, some limitations in this study have to be mentioned for future research. In regard to the experimental protocol, we only implemented at most five cycles of I/R, so the longer effect of I/R in PU remains unknown. Furthermore, as the mechanisms of I/R injury and the signaling pathway of apoptosis in the early-stage PU development are multifactorial, additional research is needed to elucidate the sequence of intracellular events and the interactive effects between HIF-1α and apoptosis.
In summary, our results showed that a mitochondria-mediated apoptotic pathway is activated in the development of early-stage PU after suffering prolonged I/R procedures. The interaction of energy depletion, oxidative stress reaction, and inflammatory response may induce apoptosis. HIF-1α plays a potential role for promoting apoptosis, which is not consistent with our original hypothesis. As apoptosis is a highly controllable, programmed biological process, one can hope to modulate or even reverse the progress of pressure ulceration by steering the apoptotic signaling pathway and regulating the HIF-1α role of cytoprotection or proapoptosis. Thus, we expect that muscle cell apoptotic signaling events and the HIF-1α gene can be the potential targeting points for PU intervention.
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