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J Appl Physiol 100: 1946-1954, 2006. First published February 16, 2006; doi:10.1152/japplphysiol.00889.2005
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Compression-induced deep tissue injury examined with magnetic resonance imaging and histology

A. Stekelenburg,1 C. W. J. Oomens,1 G. J. Strijkers,2 K. Nicolay,2 and D. L. Bader1,3

1Department of Materials Technology; 2Biomedical NMR, Eindhoven University of Technology, Eindhoven, The Netherlands; and 3Department of Engineering and IRC in Biomedical Materials, Queen Mary, University of London, London, United Kingdom

Submitted 22 July 2005 ; accepted in final form 14 February 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGMENTS
 REFERENCES
 
The underlying mechanisms leading to deep tissue injury after sustained compressive loading are not well understood. It is hypothesized that initial damage to muscle fibers is induced mechanically by local excessive deformation. Therefore, in this study, an animal model was used to study early damage after compressive loading to elucidate on the damage mechanisms leading to deep pressure ulcers. The tibialis anterior of Brown-Norway rats was loaded for 2 h by means of an indenter. Experiments were performed in a magnetic resonance (MR)-compatible loading device. Muscle tissue was evaluated with transverse relaxation time (T2)-weighted MRI both during loading and up to 20 h after load removal. In addition, a detailed examination of the histopathology was performed at several time points (1, 4, and 20 h) after unloading. Results demonstrated that, immediately after unloading, T2-weighted MR images showed localized areas with increased signal intensity. Histological examination at 1 and 4 h after unloading showed large necrotic regions with complete disorganization of the internal structure of the muscle fibers. Hypercontraction zones were found bilateral to the necrotic zone. Twenty hours after unloading, an extensive inflammatory response was observed. The proposed relevance of large deformation was demonstrated by the location of damage indicated by T2-weighted MRI and the histological appearance of the compressed tissues. Differences in damage development distal and proximal to the indenter position suggested a contribution of perfusion status in the measured tissue changes that, however, appeared be to reversible.

deep pressure ulcers; decubitus; animal model

PRESSURE ULCERS ARE AREAS of soft tissue breakdown that result from sustained mechanical loading of skin and underlying tissues. They can interfere with quality of life, activities of daily living, and rehabilitation and, in some cases, may prove life threatening. Pressure ulcers can develop either superficially or deep within the tissues, depending on the nature of the surface loading and the tissue integrity (7, 12). The superficial type forms within the skin, with maceration and detachment of superficial skin layers. If allowed to progress, the damage may form an ulcer, which is easily detected. By contrast, deep ulcers arise in muscle layers covering bony prominences and are mainly caused by sustained compression of the tissues. These ulcers develop at a faster rate than superficial ulcers and yield more extensive ulceration with an uncertain prognosis. The underlying mechanisms that lead to these deep ulcers are not fully understood. The focus of the present study is, therefore, on deep pressure ulcers, which necessarily involve deep tissue injury, recently defined as "A pressure-related injury to subcutaneous tissues under intact skin" (18). This definition instantly reveals one of the major problems associated with their early detection.

One of the populations susceptible to the development of deep pressure ulcers are spinal cord injury (SCI) subjects (14, 25). As an example, the study by Garber and Rintala (25) reported an incidence rate of 39% in a 3-yr period. A significant proportion of these were stage IV ulcers (20), two-thirds of which were associated with the pelvic region. Particular risk factors for SCI subjects include limited activity and mobility levels and sensory deficit.

To learn more about the underlying mechanisms leading to pressure ulcers, different animal models have been used over many decades. Several theories have been proposed. The most commonly adhered theory is that compression of the tissues causes occlusion of capillary blood flow, resulting in local ischemia and a depletion of the supply of vital nutrients to the cells (3, 16, 17). More recently, reperfusion following an ischemic period has been suggested as an additional factor in damage development (31). A further theory focuses on the role of the interstitium between cells. This theory assumes that mechanical loading results in a disturbance in the metabolic equilibrium around cells (40, 47). The prolonged deformation of cells, per se, has also been proposed to play a major role in the onset of tissue damage (9, 11, 13, 49).

In association with the latter hypothesis, the larger deformations found in muscle tissues of SCI subjects might be an additional risk factor for this subject group (45, 48). In a human MRI study, the total compression of the soft tissue composite above a bony prominence was estimated to be 30% for normal subjects compared with 50% for a SCI subject with flaccid paralysis (48). These values, determined with supine individuals, might be enhanced in appropriate tissues in seated individuals. Indeed, such deformations can be associated with significantly increased interface pressures in SCI subjects with flaccid paraplegics compared with other groups, such as the elderly and spastic paraplegics (45).

There has been considerable research associated with muscle damage. Most of this research has involved exercise-induced damage (37, 41), muscle diseases (29), and ischemia-reperfusion injury (22, 30). By contrast, only a few studies have examined muscle damage directly related to pressure ulcers. These latter studies have generally used histology to assess muscle damage in the form of loss of cross striation and infiltration of inflammatory cells (9, 38, 51). A more detailed examination of the histopathology, however, might reveal information that is vital to the understanding of the primary damage mechanism. In addition, information on the spatial and temporal development of damage in muscle tissue in vivo can be provided by MRI, which is a nondestructive technique.

Bosboom et al. (9) developed an animal model in which the tibialis anterior (TA) of Brown-Norway rats was compressed by means of an indenter. Muscle tissue was examined after 24 h using both histological analysis and MRI. It was shown that affected tissue localized by T2-weighted MRI correlated well with damaged areas determined by histological examination. An increase in the transverse relaxation time (T2) is generally accepted as a measure of tissue damage (21). The pathological features associated with pressure ulcers (32, 34, 54) that may affect muscle proton density and relaxation times are inflammation, edema, necrosis, hemorrhage, fibrosis, and fatty infiltration. Of these, inflammation, edema, and also hemorrhage can lead to an increased proton density, caused by an increased in both intracellular and extracellular free water. This will result in an increase in T1 (longitudinal relaxation time) and T2 because free water has longer relaxation times. T1 reflects structural aspects but is relatively insensitive to changes in the state of the muscle. In contrast, T2 is very sensitive to tissue changes.

Our hypothesis is that initial damage to the muscle fibers is induced mechanically by local excessive deformation and subsequent disruption of muscle fibers. This damage is followed by a cascade of processes as the tissue homeostasis is disturbed. These include a change in calcium homeostasis due to membrane disruption, resulting in calcium-activated degradative processes (2, 4) and postinjury inflammation that further degrades the tissue. Based on the notion that the initial muscle damage might reveal the proposed mechanical damage, the focus of the present study was to examine the early damage in muscle tissue after compressive loading. It employed a magnetic resonance (MR)-compatible loading device (53), which enabled the simultaneous application of pressure to the TA and measurement of tissue status by means of MRI. T2-weighted MRI was performed both during loading and up to 20 h after unloading, and histological examination was performed at early time points after unloading. These measurements were combined to elucidate the damage mechanisms leading to deep pressure ulcers.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 ACKNOWLEDGMENTS
 REFERENCES
 
Animal model.   In this study, 20-wk-old female Brown-Norway rats weighing between 170 and 200 g were used. They were housed under standard laboratory conditions (12-h light, 12-h dark cycles) and maintained on standard laboratory food and water ad libitum. Each rat was anesthetized for the preparation phase by subcutaneous injection of xylazine (1 µl/g body wt, 2 g/l) and intramuscular injection of ketamine (0.8 µl/g body wt, 100 g/l). During the MR measurements, anesthesia was maintained with isoflurane inhalation [0.4–1.0% isoflurane with N2O-O2 mixture (1:1)]. Vital signs (pulse and respiratory rate) were monitored and maintained within physiological values. Each rat was placed on a heating pad to maintain body temperature between 35 and 37°C. Before the loading experiment, the hairs on the left TA region were removed by shaving. The leg was placed in a specially designed mould, and a plaster cast was applied to obtain a firm fixation in the setup. The preparation phase took ~45 min. The experimental protocol was approved by the Animal Care Committee of the University of Maastricht.

MR-compatible loading device.   The experimental setup is described in detail elsewhere (53). To review briefly, the loading device consisted of two concentric tubes, the inner of which houses the animal, whereas the outer tube was used to position the whole arrangement in the MR scanner. The scanner was a 6.3-T Varian system, operating at 270 MHz (horizontal bore, diameter of 95 mm) with a 380 mT/m gradient coil. The anesthetized animal was placed supine in the loading device (Fig. 1), with its foot positioned in a special holder in which the casted leg was fixed. In the cast, a hole was made for the application of a rounded plastic indenter, of 3-mm diameter, to the TA region (Fig. 1B). A birdcage radio-frequency coil was placed in a fixed position around the limb. The indenter was fixed to a glass fiber reinforced polymer (Ertalon 66GF30) loading beam to which strain gauges were attached to enable force measurements during indentation. An indentation was applied by rotating the bar that is attached to a cam, as shown in Fig. 1A. Figure 1C shows a transversal MR image of the lower limb underneath the indenter during indentation.


Figure 1
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  		Fig. 1. A: schematic of magnetic resonance (MR)-compatible loading device. Animal is lying supine in the setup with its leg fixed in the radio-frequency (RF) coil. B: photograph of an expanded view of the loading device in which pressure is applied to the tibialis anterior (TA) by an indenter (i) that is positioned through the legs of the RF coil. C: transversal MR image showing the TA (indicated by white line) compressed between tibia and indenter. The indenter was filled with water to ensure visibility on the MR image.

 
Experimental protocol.   Animals (n = 10) were divided into three groups (designated group I to group III). The loading protocol was the same for all groups, whereas the moment of perfusion fixation was different.

The measurement protocol involved five separate phases, as indicated in Fig. 2. Initially transversal scout images were produced, and, where necessary, adjustments were made to ensure that the TA was compressed between the indenter and tibia and that the indentation was perpendicular to the surface of the limb. Before the indenter was applied, a T2 map (T2p) was measured using a multi-echo sequence over a scanning time of 20 min. Thereafter, an indentation of 4.5 mm was applied, at a rate of 1.5 mm/s to avoid impact damage, and maintained for 2 h. During indentation, sagittal and coronal images and a T2 map (T2L) of the leg were recorded. Third, immediately after removal of the indenter, a series of T2 measurements was started (T21–8). For group I animals (n = 3), two measurements were performed in a 1-h period, after which the animal was killed for histological examination. For group II animals (n = 4), up to eight consecutive measurements were performed, and at 4 h the animals were killed. After completion of the measurements, group III animals (n = 3) were removed from the loading device, their cast was removed, and they were allowed to recover from anesthesia and move freely for the next 16 h. After this period, the animal was again sedated and the leg fixed in the setup. A single T2 measurement (T29) was performed after which the animal was killed (Fig. 2).


Figure 2
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  		Fig. 2. Time schedule of MR measurements and death for histological examination. Animals (n = 10) were subjected to one loading protocol and were killed for histological examination at 3 different time points, designated groups I, II, and III, respectively. Shaded rectangle represents loading period. sc, Scout images; T2, transverse relaxation time (T2) map; sag/cor, sagittal and coronal images; H, histological analysis; subscripts p, L, and 1–9, preloading, during loading, and number of measurements after unloading, respectively; t = 0, moment of unloading.

 
MRI parameters.   A multi-echo spin echo sequence was used to obtain T2 maps. Signal intensities of the echoes were fitted to the equation SI = S0 exp (–t/T2) + C, where SI is signal intensity, S0 is the intensity at 0 ms, t is time, and C is a constant to determine T2 values. Imaging parameters were slice thickness = 1 mm, field of view = 25 x 25 mm2, matrix size = 128 x 128 pixels, number of signal averages = 2, echo time = 12–96 ms, number of echoes = 8, and repetition time = 4.5 s. Number of slices = 41, centered on the longitudinal axis of the indenter. Total scanning time was 20 min.

A T2-weighted spin echo sequence was used to collect 31 slices in the sagittal and coronal direction, perpendicular to the indenter direction. Imaging parameters were slice thickness = 1 mm, field of view = 60 x 30 mm2, matrix size = 128 x 128 pixels, number of signal averages = 2, echo time = 25 ms, and repetition time = 4 s. Total scanning time was 17 min.

MR data analysis.   To obtain an enhanced contrast-to-noise ratio between normal and affected tissue, the individual images of the multi-echo sequence were summed. The resulting images are referred to as T2-weighted-sum images. On the T2-weighted-sum images that were collected preloading (T2p), the TA region was manually defined, which excluded both the skin and fat layer. In this area, a mean T2 value was calculated, and a threshold value was defined as equal to the mean + 2 SD. This threshold was applied to all images up to T28 (Fig. 2). The affected area was then defined as the percentage number of pixels with signal intensity above the threshold value. Spatial and temporal development of affected regions was evaluated in volumes consisting of two adjacent 1-mm-thick slices. Three volumes were used for evaluation: underneath the indenter and at 3 mm proximal and distal to the indenter. For the calculation of temporal development of damage, only those experiments were included in which eight consecutive T2 maps had been collected (n = 4). In three experiments, MR measurements were missing due to scanner problems or breathing instabilities of the animal. Data are presented as means ± SE. For images of group III animals taken on day 2, slices were retraced according to the size and shape of the tibia bone.

Statistical analysis.   SPSS version 12.0.1 (SPSS, Chicago, IL) was used for statistical analysis. Comparison between distributions of T2 values was performed using a nonparametric test (Mann-Whitney). A two-tailed paired Student's t-test was performed to test differences in temporal evolution and to test differences between volumes distal and proximal to the indenter. In all cases, P < 0.05 was considered significant.

For statistical analysis of MR measurements, experiments from groups II and III were used. All animals were subjected to the same loading protocol. An estimated value for {sigma}/{delta} of 0.5, a power level of 0.8, and a significance level {alpha} = 0.05 indicated a necessary group size for analysis of n ≥ 4.

Histological analysis.   Immediately following the last MR measurement, each animal was killed and perfusion fixated with 4% buffered formalin. The lower limb was cut off and stored in formalin. The TA was excised at least 2 wk later to ensure complete tissue fixation. The muscle was dehydrated in a series of alcohol solutions and embedded in plastic (Technovit 7100, Kulzer). The muscle was cut longitudinally, perpendicular to the direction of load application, or in the transversal direction in 5-µm-thick sections. The samples were stained with Gomori's trichrome to visualize both the cross-striated appearance of the muscle fibers and the cell nuclei.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 ACKNOWLEDGMENTS
 REFERENCES
 
MRI.   Figure 3 illustrates transversal T2-weighted-sum images of the lower limb underneath the indenter recorded before, during, and after loading. The image taken during loading (T2L; Fig. 3B) was collected just before load removal and revealed no systematic changes in signal intensity in the loaded region. In the slices distal to the indenter position, some signal increase was observed during loading. Figure 3C shows the image taken directly after unloading (T21) and clearly demonstrates a marked signal increase in the TA region in a localized area extending from skin down to the tibial bone. There was considerable similarity between all experiments in locations of the areas exhibiting an increased signal intensity.


Figure 3
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  		Fig. 3. Transversal T2-weighted-sum images taken before loading (T2p; A), during loading (T2L; B), and immediately after unloading (T21; C). TA is indicated by the white line in A.

 
A typical temporal distribution of T2-values in the TA region underneath the indenter is shown in Fig. 4. The T2 distribution before loading appeared to be symmetrical (skewness 0.236) with a mean value of 20.3 ± 2.7 ms (Fig. 4A) and a threshold value of 25.7 ms, which is indicated by the vertical line. Pixels with a T2 above this value are considered to be indicative of muscle damage. The distributions associated with the postloading periods were clearly skewed toward higher T2 values (skewness 0.802, 1.479, 1.419, respectively). Thus the median T2 value was 26.8 ms at 15 min and had increased to 30.0 ms at 90 min after unloading, an increase that was statistically significant (P < 0.001). Thereafter, there was a statistically significant decrease (P < 0.001) to a value of 27.9 ms corresponding to 190 min after load removal.


Figure 4
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  		Fig. 4. Distribution of T2 values in TA underneath the indenter. A: before loading, the distribution is symmetrical with a mean value of 20.3 ± 2.7 ms. Threshold value (mean + 2 SD) is indicated by vertical line. After load removal, a skewed distribution toward higher T2 values was found with a median of 26.8 ms at 15 min (B), 30.0 ms at 90 min (C), and 27.9 ms at 190 min (D) after load removal.

 
The trends in temporal response are more closely illustrated in Fig. 5. The percentage number of pixels with a T2 above the threshold value are shown for 15–190 min after load removal. Trends are illustrated for three volumes (Fig. 5A), underneath the indenter and at 3 mm proximal and 3 mm distal to the indenter position. It should be noted that the largest percent affected volume was found underneath the indenter (volume 2 in Fig. 5A). This volume slightly increased in the first 90 min to 60%, after which it tended to decrease. No significant changes occurred in the first 3 h after unloading. In both volumes at 3-mm distance from the indenter, the affected percentages were significantly lower than in volume 2 (P < 0.001 at all time points). Two different trends were observed. In the slices proximal to the indenter (volume 1 in Fig. 5A), only a small affected volume was evident directly after unloading, which increased in the first 90 min from 11 to 33%, the increase being statistically significant (P < 0.05). Thereafter, it tended to decrease. In the slices distal to the indenter, the percent number of affected pixels was 39% at 15 min, which decreased to 25% in the first hour after unloading.


Figure 5
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  		Fig. 5. Temporal and spatial distribution of affected tissue characterized by a significantly prolonged T2. A: sagittal slice of lower limb with 3 volumes selected for analysis proximal to (1), underneath (2), and distal to (3) the indenter. B: percentage number of pixels in selected volume above threshold value as a function of time. Percents are indicated as means ± SE (n = 4).

 
The T2 maps underneath the indenter recorded before (T2p) and during loading (T2L), and 190 min (T28) and 20 h (T29) after unloading are shown in Fig. 6. The regions in the TA with enhanced signal intensity exhibited a similar distribution at 20 h compared with 190 min after unloading for all three experiments (group III animals). The high T2 values evident at the edges (Fig. 6, B and C) were caused by water accumulation in the skin layer.


Figure 6
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  		Fig. 6. T2 maps of lower limb underneath the indenter taken before (A) and during loading (B), and at 190 min (C) and 20 h (D) after unloading. The different shape of the leg in D is caused by small differences in casting between days 1 and 2.

 
Histology.   Longitudinal slices taken from muscle fixated 60 min after load removal (group I animals) are illustrated in Fig. 7, A and B. Two regions are shown taken from the midportion of the muscle, where the muscle was compressed by the indenter. A large area was evident with a complete absence of the normal cross-striated pattern of skeletal muscle, indicating a disorganization of the internal structure of muscle fibers. The filaments were "snapped" and clustered together, leaving empty spaces within the fibers. The morphology of individual fibers indicated both necrotic regions and normal appearance separated by segmental contraction. In those slightly darker stained parts of the fiber, the diameter was clearly increased (Fig. 7A, short arrows). At the interface between hypercontracted and undamaged fibers, longitudinal tears were observed (Fig. 7B, arrows). Adjacent to the necrotic region, the membrane of the fibers had a wavylike appearance and the interstitial spaces were widened. The histological slices of muscles from group II animals fixated 4 h after load removal (Fig. 7, C and D) showed necrotic areas with similar structural damage to the muscle fibers. In some fibers, the contractile material was slightly less clustered together. At the edges of the necrotic region, early infiltration of polymorphonuclear cells (PMNs) was observed (Fig. 7C, arrows). In some parts of the muscle, there was an alternating pattern of damaged and undamaged fibers. Normal cross-striated fibers were found adjacent to completely disrupted fibers (Fig. 7D). The membranes of the disrupted fibers seemed largely intact, with only minor evidence of fragmentation. After 20 h of load removal (Fig. 7, E and F), the cellular debris was more diffusely distributed within the fibers, occupying the empty spaces observed in the first hours after load removal. This cellular material was stained lighter compared with the undamaged filaments. It is interesting to note that, despite the extensive regions of necrosis, the individual fibers could clearly be distinguished. An extensive infiltration of PMNs and monocytes was observed, revealing a pronounced inflammatory response. These inflammatory cells aggregated in areas of the interstitium of the muscles and invaded a proportion of muscle fibers.


Figure 7
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  		Fig. 7. Longitudinal histological slices of TA stained with Gomori's trichrome. Muscle fixated 1 h (A and B), 4 h (C and D), or 20 h (E and F) after removal of load. The muscle fixated 1 h after load removal showed a complete disorganization of the internal structure of the muscle fibers. After 4 h (C and D), early signs of inflammation were visible by infiltration of polymorphonuclear leukocytes (C, arrows). After 20 h (E and F), an extensive infiltration of polymorphonuclear neutrophils and monocytes is visible. Bar represents 50 µm.

 
Transversal histological slices of muscle fixated 20 h after load removal are illustrated in Fig. 8. These slices more clearly demonstrate the distribution of size and shape of the muscle fibers. In undamaged regions, fibers were angular in shape and show a uniform intensity of staining. The injured region showed damage ranging from slightly different staining intensity and rounded appearance of fibers (indicated by arrows, Fig. 8C) to necrotic, pale-stained fibers (Fig. 8A). An extensive infiltration of PMNs and monocytes was also observed in these slices. Some fibers showed phagocytotic infiltration (indicated by arrow, Fig. 8B). It is also evident that these large fibers compressed surrounding fibers, indicated by asterisks in Fig. 8B. Overall, the distribution of necrotic fibers within the fascicles did not reveal a systematic pattern.


Figure 8
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  		Fig. 8. Transversal slice of TA stained with Gomori's trichrome from muscle fixated 20 h after unloading. A: mixture of damaged (necrotic) fibers (light) and undamaged fibers (dark). The infiltration of polymorphonuclear leukocytes and monocytes (dark dots) revealed an extensive inflammatory reaction. B: mononuclear cells aggregated in localized areas of the interstitium of the muscles and in some muscle fibers (arrow). The large fiber compressed surrounding fibers (*). C: round swollen cells, with normal staining intensity, lie scattered within the cross sections (arrows). Bar represents 200 µm (A), 100 µm (B), and 50 µm (C).

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
ACKNOWLEDGMENTS
 REFERENCES
 
In the present study, muscle damage after compressive loading was examined with T2-weighted MRI in combination with histological examination. Affected regions within the muscle tissue could clearly be distinguished on T2-weighted images. In addition, the subsequent temporal development of damage could be monitored. Histological examination at early time points after unloading revealed initial damage to the muscle fibers.

The correlation between MRI and histology is not straightforward as changes in T2 can reflect a range of pathologies (21). Bosboom et al. (9) examined the correlation between the increase in T2 and histological damage in rat skeletal muscle tissue at one time point. An extensive and labor-intensive method was used to correlate damage in histological slices to high intensities in T2-weighted MR images, both determined 24 h after load removal. Damage in the histological slices was indicated manually from evidence of loss of cross-striation of the muscle fibers and/or infiltration of inflammatory cells. Damage in MR slices was determined by applying a threshold level to the images. The correlation found between the damaged areas as quantified using MRI and histology, 24 h after load removal, was very high (r2 = 0.93). In the present work, the histological damage observed 20 h after load removal appeared similar to that described in the previous study. It may therefore be inferred that the increase in T2 at 20 h after unloading, as was observed in the present study, was a direct result of the change in appearance of the muscle fibers, as found in the histological slices (Fig. 7, E and F, and Fig. 8).

The correlation between T2 increase and changes in tissue integrity at earlier time points after load removal is, however, more complex in nature. In the histological slices taken 1 and 4 h after load removal several changes were observed. Primarily, in the area underneath the indenter, large necrotic regions were observed with condensed contractile material clustered together within the fibers. Adjacent to this necrotic region, there was a less systematic picture. Areas contained both necrotic fibers and fibers exhibiting different stages of damage, including longitudinal tears, wavylike membranes, probably with changed permeability, widened interstitial spaces and hypercontracted zones. These factors will probably all contribute to an increase in T2. It is important to note that the T2 value in a MRI pixel is a mean value of a region containing ~20 fibers with a length of 1 mm, equivalent to the thickness of the MR slice. Thus the resolution of MRI is considerably lower than that provided by histology. Therefore, in the present study, only a qualitative evaluation was performed. For a direct correlation between MRI and histology, the uptake of Evans blue dye, a histological marker of muscle damage, could prove useful (24, 27). In addition to providing a quantitative correlation, this approach could provide information on the status of membranes of nonnecrotic fibers.

In this study, a monoexponential T2 relaxation was assumed. This approach was adopted because the signal-to-noise ratio and the number of echoes used in the present study precluded the use of any nonmonoexponential analysis. However, several studies, using more extensive sequences, have demonstrated alternative forms of relaxation. For example, biexponential decay curves have previously been related to the intra- and extracellular water compartments (1, 15). Other studies (19, 28, 50) have demonstrated three ranges for T2 values: T2 < 5 ms for water associated with macro molecules, 25 < T2 < 45 ms for intracellular water, and T2 > 100 ms for extracellular water. More extensive measurements, revealing those different components, might add useful information in future experiments, although this approach will compromise the time resolution of the experiments.

In this study, large indentations, equivalent to surface pressures of ~150 kPa, were applied, simulating the indentation of muscle tissue by bony prominences (43, 48). Underneath the indenter, no systematic changes in signal intensity were observed during loading. Although this finding suggests that no large changes in water balance occurred in the loading phase, it does not indicate whether the tissue was affected when exposed to large deformations, since T2 does not reflect all changes to tissue and the resolution is limited. Directly after unloading, a large area with increased T2 values was present, tending to decrease in size after 2 h. A striking difference was observed between slices proximal and distal to the indenter position (Fig. 5). In the slices proximal to the indenter, T2 values started to increase after unloading. The affected volume increased in the first hour, after which it slowly decreased. In the slices distal to the indenter, however, higher T2 values were already observed during and directly after load removal. However, the affected area decreased in the first hours. The deformations in both these regions are necessarily small. Therefore, a possible explanation for the different responses is the perfusion status of the tissue. The perfusion in the tissue distal to the indenter was presumably blocked by the large indentation, whereas in the slices proximal to the indenter the perfusion was minimally affected during loading. The difference in amount of affected volume largely disappeared in the first hour, implying that the effect of ischemia, as indicated by an increase in T2, was reversible within the muscle tissue.

The large deformations underneath the indenter result in large strains on the muscle fibers. These high strains on individual muscle fibers resulting from compressive loading may be similar to those reported associated with exercise. The two common hypotheses (41) to explain exercise-induced damage are metabolic overload and mechanical factors, the latter of which may have direct relevance to the present study. Indeed, Lieber and Fridén (36) demonstrated in an in vivo model that muscle damage after eccentric contraction was a function of active muscle strain as opposed to muscle force. The possible analogy between the characteristics of damage after compression and exercise may be examined by reproducing a histological schematic of damage after eccentric exercise (Fig. 9B), indicating a muscle fiber with segmental damage surrounded by normal fibers (23). Features include hypercontraction zones, bilateral to the necrotic zone, which displace and compress adjacent fibers, whereas in the region of the lesion these normal fibers taper along the damaged and narrow fiber, all of which are illustrated in Fig. 9B. These features in damaged fibers were also found in the present histology after compressive loading as illustrated in Fig. 9A, suggesting a similar mechanical damage mechanism. Although large strains might in both damage processes be a crucial factor, the level at which the damage initiates is probably different. The damage after eccentric exercise is proposed to initiate at the sarcomere level (44, 46). The popping sarcomere hypothesis (42) states that stretch-induced muscle damage results from very nonuniform lengthening of sarcomeres when active muscle is stretched beyond optimum length. This nonuniform lengthening leads to shearing of myofibrils, exposing membranes to large deformations. This leads to loss of calcium ion homeostasis. Compared with exercise, the large deformations applied to the tissue in the present study represent a more aggressive insult to the muscle. An early necrotic response was observed after unloading. The large necrotic region at the position of indentation is assumed to be caused by the large deformation that simply pulled the muscle fibers apart. In the regions near the indenter, the extreme axial strain of the membrane is proposed to create invaginations and tearing that opens calcium channels, which also leads to loss of calcium ion homeostasis. Disrupted calcium homeostasis can, in both cases, result in calcium-activated degradative processes (2, 4).


Figure 9
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  		Fig. 9. Segmental damage to muscle fiber. A: longitudinal slice of muscle fixated 1 h after a 2-h loading period, stained with Gomori's trichrome. Hypercontraction zones (arrow) are bilateral to the necrotic zone. Bar represents 50 µm. B: schematic drawing of longitudinal section of muscle after eccentric exercise showing the different zones, normal-hypercontraction (arrow)-necrotic (middle)-hypercontraction (arrow)-normal, in a fiber with segmental damage. Adapted from Fridén and Lieber (23).

 
Different studies on the effect of stretching muscles (6, 13a, 36) have shown the importance of strain on muscle fibers as an initial damage mechanism. Local strains might be correlated to damage location by using MR-tagging experiments of local tissue deformation, involving separate repetitive indentation phases. Another approach to estimate strains during indentation is to develop a dedicated finite element model to examine the spatial relationship between damage and predicted strains. Both methods will be adopted in future studies.

It is important to stress that the number of animals participating in this study was limited. The used MR-compatible loading device, however, allowed the exact measurement of the applied indentation, contrary to previous studies, in which only the applied pressure could be measured. Therefore, the similarity between all experiments was considerable. In view of the mechanical hypothesis, the exact amount and location of damage will depend on the local strain fields. These will be different for each experiment, despite the controlled load application. This supports the proposed approach in which each individual loading arrangement is modeled using a dedicated finite element model.

The large deformations used in this study are considered to be relevant, especially for SCI subjects. It is known that muscle tissue properties change after SCI. This will inevitably have an influence on the local deformation of muscle tissue. In a review by Scelsi (52), the pathological changes were summarized and included muscle fiber-type transformation, with type I fiber change to type II, muscle atrophy, changes in fiber size, and alterations in myofibrillar apparatus. Studies on an experimental spinal cord transection showed changes in rat skeletal muscle, with almost complete type I to type II fiber transformation after 1 yr (35). The use of such an animal model might be valuable in studying the vulnerability of SCI subjects. Several pressure ulcer-related studies involving paraplegic animals have been performed (16, 26, 33). Results were, however, contradictory, caused, in part, by differences in time between spinal cord transection and experiments, which seemed to represent a critical factor.

It is well accepted that MRI will not be available or even practical for use with all patients at risk. Thus a prescreening method is required, incorporating the identification of physical markers and early damage markers, which can be measures in, for example, blood. These markers can be used as a prescreening indicator for subsequent MR scanning.

In summary, the proposed relevance of large deformation of muscle cells in the development of deep tissue injury was supported by the location of damage indicated by T2-weighted MRI and a resemblance in histological appearance between exercise-induced and compression-induced muscle damage. Based on the difference in the response of muscle tissue observed distal and proximal the indenter, which was most likely an effect of perfusion, the increase in T2 revealed tissue changes due to ischemia alone. These tissue changes will most probably accelerate and/or increase the damage development during and after compressive loading.


   ACKNOWLEDGMENTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGMENTS
REFERENCES
 
We gratefully acknowledge Jo Habets for help with the animal experiments and Dr. F. Verheyen from the University of Maastricht and Dr. M. Lammens from Radboud University Nijmegen Medical Centre for help with the interpretation of the histological slices.


   FOOTNOTES
 

Address for reprint requests and other correspondence: A. Stekelenburg, Eindhoven Univ. of Technology, Dept. of Biomedical Engineering, PO Box 513, Den Dolech 2, 5600 MB Eindhoven, The Netherlands (e-mail: A.Stekelenburg{at}tue.nl)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGMENTS
 REFERENCES
 

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