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In many muscle pathologies, impairment of skeletal muscle function is closely linked to changes in the mechanical properties of the muscle constituents. Optical coherence micro-elastography (OCME) uses optical coherence tomography (OCT) imaging of tissue under a quasi-static, compressive mechanical load to map variations in tissue mechanical properties on the micro-scale. We present the first study of OCME on skeletal muscle tissue. We show that this technique can resolve features of muscle tissue including fibers, fascicles and tendon, and can also detect necrotic lesions in skeletal muscle from the mdx mouse model of Duchenne muscular dystrophy. In many instances, OCME provides better or additional contrast complementary to that provided by OCT. These results suggest that OCME could provide new understanding and opportunity for assessment of skeletal muscle pathologies.
© 2014 Optical Society of America
1. Introduction
The mechanical properties of skeletal muscle tissue, from the sub-cellular to the organ level, are integral to muscle function [1]. During normal skeletal muscle activation, contractile forces are generated by actin/myosin protein filaments within individual muscle cells. These sub-cellular forces are transmitted across the cell membrane (sarcolemma) to the extracellular matrix, and through connective tissue and tendons [2]. The structure and mechanical properties of these muscle constituents govern the aggregate skeletal muscle behavior. By studying the mechanical behavior of skeletal muscle, it is possible to infer the function of muscle constituents, for example, the molecular mechanisms of muscle contraction can be probed through examination of the mechanical properties of activated skeletal muscle [3]. In many pathologies, such as Duchenne muscular dystrophy (DMD), impairment of muscle function is associated with changes in the mechanical properties of the muscle constituents [4, 5]. DMD is a progressive, lethal, muscle-wasting disorder affecting 1 in 3600 to 6000 live male births [6], and caused by defects in the gene which codes for the sub-sarcolemmal protein dystrophin [7]. The lack, or impaired functioning, of dystrophin leads to changes in the mechanical properties of muscle tissue on the cellular level: dystrophin-negative muscle fibers are highly susceptible to contractile damage, activating an inflammatory response which leads to necrosis of the muscle fibers [8]. In DMD patients, repeated cycles of necrosis and regeneration leads to the formation of regions of necrotic muscle fibers, inflammatory cells, and fibrous and fatty tissue, referred to as necrotic lesions [9], eventually resulting in a loss of muscle function at the macro-scale.
Changes in the mechanical properties of skeletal muscle at the cellular and sub-cellular levels can be detected using methods such as atomic force microscopy and micro-pipette aspiration [10, 11]. Whilst these techniques allow skeletal muscle cell mechanics to be probed, generally they can only be performed on excised tissue. Changes in the mechanical properties of skeletal muscle at the macro-scale can be measured in vivo with medical elastography techniques based on ultrasound or magnetic resonance imaging (MRI) [12, 13]. However, the initial formation of necrotic lesions occurs on the intermediate scale, between that of cells and whole organs [9, 14, 15]. The limited spatial resolution of medical elastography techniques, and the limited field-of-view and depth penetration of cellular imaging modalities, makes them both ill-suited for imaging mechanical contrast on this scale. A method for investigating mechanical properties on an intermediate scale could provide new information to advance the understanding and treatment of muscle pathologies such as DMD.
One such emerging method is optical coherence elastography (OCE) [16], which utilizes optical coherence tomography (OCT) to measure tissue deformation in response to static or dynamic loading [17, 18]. There has been much progress in OCE, with many different loading mechanisms and imaging techniques under development [19–24]. Recently, our group has demonstrated an OCE technique, dubbed optical coherence micro-elastography (OCME), which improves upon phase-sensitive compression OCE to enable high-resolution imaging (close to the OCT lateral resolution) of mechanical contrast in tissue over relatively large (~1 × 1 cm) en face fields of view. We have shown the ability of this method to provide micro-scale mechanical contrast in human breast and lymphoid tissue [25]. Other studies on OCE have proposed a number of applications, including in ophthalmology [26–28], cardiology [29] and dermatology [30–32]. Some recent studies have demonstrated OCE’s potential for imaging tendon [33] and cardiac muscle [34]; however, no dedicated study has been performed to investigate OCE’s potential for imaging skeletal muscle.
In this study, we demonstrate the ability of OCME to provide mechanical contrast in skeletal muscle tissue from the mdx mutant mouse strain (C57BL/10ScSnmdx/mdx), which is widely used in pre-clinical research as an animal model for DMD [35]. We show that OCME can image the entire surface of mouse skeletal muscles, with micro-scale resolution sufficient to detect individual muscle fibers, fascicles, and tendons, and also to detect areas of necrosis in mdx skeletal muscle tissue, as validated by histology. Previous studies have examined the use of OCT as a tool for assessing skeletal muscle pathologies [36–40]. Our results in this study demonstrate that OCME provides contrast, in many instances, better than or additional to the corresponding OCT.
2. Materials and methods
2.1. Optical coherence micro-elastography imaging
The OCME system used in this study has been previously described in detail [25], and is briefly described here. The system is based on Fourier-domain OCT [41] employing a superluminescent diode light source with a mean wavelength (λ0) of 835 nm and a 3-dB spectral bandwidth (Δλ) of 50 nm, which illuminates the sample with 10 mW of optical power. The measured free-space axial/lateral resolution (full-width at half-maximum irradiance) is 8.5 µm/11 µm. The interference spectrum for each A-scan is captured over 1,300 pixels of a CMOS line-scan camera with an exposure time of 2 µs. The system operates in a common-path configuration, with the reflection from the imaging window-tissue interface acting as the reference reflector.
The imaging window (thickness, 2 mm) is fixed to a piezo-electric ring actuator, enabling mechanical loading and imaging of the sample from the same side [25, 41, 42]. A rigid brass plate, of larger surface area than the sample, is used to apply a preload to ensure uniform contact between the plate, the sample, and the imaging window. The ring actuator is driven by a 5-Hz square wave synchronized with the B-scan acquisition frequency of 10 Hz, so that consecutive B-scans are acquired in the unloaded/loaded state [25, 41]. A schematic of the system is shown in Fig. 1.
Fig. 1 (a) Schematic of our fiber-based OCME system. SLD: superluminescent diode; PC: polarization controller; FC: fiber coupler. (b) Magnified schematic of the OCME sample stage, corresponding to the red-dashed box in (a). The top of the scan volume (zmin) is adjacent to the imaging window; the bottom of the scan volume (zmax) is located close to the rigid plate.
In this study, three-dimensional (3-D) OCME scans were acquired with dimensions (x × y × z) 10 × 10 × 3.1 mm. At each lateral (x, y) location, the depth-resolved axial displacement of the sample in response to the load was calculated from the phase difference between corresponding OCT A-scans in consecutive unloaded/loaded B-scans [43]. The scans were oversampled in the y-dimension to provide phase-correlation between sequential B-scans, which were acquired 1 µm apart [25]. The phase difference was additionally averaged over 5 pairs of OCT B-scans, and subject to phase unwrapping [25]. The local axial strain was then calculated using weighted-least-squares (WLS) linear regression over a 100 µm sliding window on the axial displacement, with the weights determined by the OCT signal-to-noise ratio (SNR) corresponding to each displacement measurement [44]. The 3-D strain volumes were then digitally sliced to produce images (micro-elastograms) indicative of the micro-scale mechanical properties of the samples.
2.2. Animal handling, sampling and histology
Three dystrophic mdx (C57BL/10ScSnmdx/mdx) and three control wild-type C57 (C57BL/10ScSn) male, twelve-week-old mice were obtained from the Animal Resources Centre, Murdoch, Western Australia. All experiments conformed to the guidelines of the National Health & Medical Research Council Code of Practice for the Care and Use of Animals for Scientific Purposes (2004) and the Animal Welfare Act of Western Australia (2002) and were approved by the Animal Ethics Committee of The University of Western Australia (RA/3/100/1063).
Two of the mdx mice completed an exercise protocol consisting of horizontal treadmill running at 12 m/min for 30 min to increase the amount of skeletal muscle damage so that, by 48 hours after the exercise, ~10–15% of the muscle was necrotic [45]. Immediately after exercise, the two mdx mice were injected with 1% (wt/vol) Evans blue dye (EBD; Sigma, St. Louis, Missouri) in phosphate-buffered saline (PBS) (pH 7.5) by intraperitoneal injection at a dose of 100 µL per 10 g body weight [35]. Our previous studies have shown that EBD is a suitable in vivo marker to guide OCT imaging of muscle to areas of leaky or necrotic myofibers [37, 38, 40]. All mice were anaesthetized with 2% vol/vol isoflurane (Bomac, Australia), N2O and O2, and euthanized via cervical dislocation. In the case of the two treadmill-exercised mdx mice, this took place 48 hours after EBD injection. Skeletal muscles from the forelimbs (triceps) and hindlimbs (gastrocnemius, quadriceps and gluteus) were excised and placed into PBS on ice for transport and subsequent imaging, which occurred within six hours of excision. A total of 32 muscles from the six mice were used in this study.
Immediately after imaging, samples were pinned to cork boards, or placed into histology cassettes, to preserve their orientation, before being fixed in neutral buffered formalin (Confix Clear pH 7.0, AB1020.1000C; Australian Biostain). Samples were then embedded in paraffin wax blocks, and sectioned into 5 µm slices, with 100 µm between adjacent sections, and stained with either hematoxylin and eosin (H&E), or Masson’s trichrome. Areas of necrosis are readily visible in both stains. Masson’s trichrome additionally stains for connective tissue and tendons, and was used on several samples to facilitate identification of tendon.
3. Results
In this section, we show representative micro-elastograms with comparison to co-located OCT images and histology from five samples: the left gluteus from a C57 mouse, the right quadriceps from a C57 mouse, the left gluteus from a treadmill-exercised mdx mouse, and the left gastrocnemius and right gluteus from a non-exercised mdx mouse. The OCT images (SNR of irradiance presented on a dB scale) and micro-elastograms (presented on a milli-strain scale) are representative en face slices taken from the 3-D data sets; the OCT images were taken from the top of the fitting range of the corresponding micro-elastograms. In this orientation, the micro-elastogram lateral resolution is isotropic and matched to the OCT lateral resolution [25]. For the magnified regions shown in Figs. 2–6, red and yellow borders indicate regions of undamaged muscle tissue, green indicates tendon, and blue indicates regions with necrosis. In Section 3.4, we present a detailed analysis of the contrast provided by OCME in murine skeletal muscle, using data from gluteus and gastrocnemius muscles from two C57 mice.
Fig. 2 Images of the right gluteus muscle from a C57 mouse. En face (a) H&E-stained histology section, (b) OCT image and (c) micro-elastogram. (d)–(i) Magnified insets (2 × 2 mm) highlighting the longitudinal striation patterns characteristic of corresponding regions of undamaged muscle fibers. OCT images and micro-elastograms were taken 100 µm from the surface (Media 1, size 10 × 10 × 1.8 mm (x × y × z)).
3.1. C57, healthy wild-type mice: gluteus and quadriceps
Figure 2 shows the results of OCME applied to the excised left gluteus muscle of a C57 wild-type mouse. The muscle fibers stain pink in the H&E histology (Fig. 2(a)), and the magnified insets (Figs. 2(d) and 2(e)) show the intact fibers with peripheral nuclei characteristic of healthy skeletal muscle. The muscle fibers are visible in the OCT images (Figs. 2(b), 2(f) and 2(g)), forming characteristic longitudinal striation patterns consistent with the parallel arrangement of individual muscle fibers [37, 39]. Similar striations are visible in the micro-elastograms (Figs. 2(c), 2(h) and 2(i)), with muscle fibers typically exhibiting less strain than surrounding tissue. In muscle physiology, the term “striation patterns” typically refers to the transverse dark and light bands caused by the actin/myosin filaments of the sarcomere; however, in this paper we use the term to refer to the light and dark longitudinal bands in the OCT images and micro-elastograms of muscle fibers. Notably, the striations in the micro-elastograms generally show higher contrast than the striations in the OCT images. A 3-D visualization of the micro-elastogram volume is presented in Media 1, showing striations robustly visualized over a range of depths, although estimates of strain become noisy at greater depths due to the decrease in OCT SNR, as expected [25, 44]. In Figs. 2(d)–2(i), magnified regions corresponding to the red and yellow boxes in Figs. 2(a)–2(c) highlight the longitudinal striation patterns of muscle fibers. The spacing between the striations is consistent with the expected size of individual muscle fibers (20–100 µm in diameter [1, 35]); this is analyzed in greater detail in Section 3.4.
Figure 3 shows the results of OCME applied to the excised right quadriceps muscle of a C57 wild-type mouse. Masson’s trichrome stains the muscle fibers red and connective tissues blue-green in the histology section (Figs. 3(a), 3(d) and 3(e)). This stain is particularly useful in readily identifying the main tendon connected to the quadriceps muscle (Fig. 3(e)). As in Fig. 2, muscle fibers are visible by the longitudinal striations in the OCT images (Figs. 3(b) and 3(f)) and micro-elastograms (Figs. 3(c) and 3(h)). The contrast of the striation patterns, and hence the orientation of the muscle fibers, is more clearly visible in the micro-elastograms than in the OCT images. The large tendon appears in the OCT image (Fig. 3(g)) and micro-elastogram (Fig. 3(i)) as a region of relatively uniform OCT SNR and strain, respectively, but the contrast between muscle fibers and tendon is more pronounced in the micro-elastograms.
Fig. 3 Images of the right quadriceps muscle from a C57 mouse. En face (a) Masson’s trichrome-stained histology section, (b) OCT image and (c) micro-elastogram. (d), (f) and (h) Magnified insets (2 × 2 mm) of corresponding regions of undamaged muscle fibers. (e), (g) and (i) Magnified insets of corresponding regions of a large tendon. OCT images and micro-elastograms were taken 100 µm from the surface.
3.2. mdx, exercised, severe pathology: gluteus
Figure 4 shows the results of OCME applied to the left gluteus muscle from a treadmill-exercised mdx mouse. The exercise protocol greatly elevated the levels of necrosis in this muscle sample; a large necrotic lesion is visible in the H&E-stained histology (Figs. 4(a), 4(d) and 4(e)) as an area with a high density of small purple-stained inflammatory cells and damaged muscle fibers. In the OCT images, the same region appears as an area which lacks the striation pattern of intact muscle fibers (Figs. 4(b), 4(f) and 4(g)), in agreement with previous studies on OCT imaging of muscular dystrophy [37]. The micro-elastograms also display a lack of striations in the necrotic lesion, but additionally possess a slightly mottled appearance (Figs. 4(c), 4(h) and 4(i)), potentially because of heterogeneities in localized muscle damage. A 3-D visualization of the elastogram data (Media 2) shows that this mottled pattern is consistently visualized over a range of depths in the necrotic region.
Fig. 4 Images of the left gluteus muscle from a treadmill-exercised mdx mouse. En face (a) H&E-stained histology section, (b) OCT image and (c) micro-elastogram. (d)–(i) Magnified insets (2 × 2 mm) of corresponding regions of a large necrotic lesion. OCT images and micro-elastograms were taken 100 µm from the surface (Media 2, size 10 × 10 × 1.8 mm (x × y × z)).
3.3. mdx, unexercised, mild pathology: gastrocnemius and gluteus
Figure 5 shows the results of OCME applied to the excised left gastrocnemius muscle from an mdx mouse. This sample was taken from the mdx mouse that did not undergo the treadmill exercise protocol; however, even without specific exercise, the skeletal muscles of mdx mice possess a low number (< 5%) of necrotic fibers [46]. Similarly to Fig. 2, areas of undamaged muscle appear in the histology as regions with intact fibers and few inflammatory cells (Figs. 5(a) and 5(d)). In the OCT images (Figs. 5(b) and 5(f)) and micro-elastograms (Figs. 5(c) and 5(h)), these areas possess the characteristic longitudinal striation patterns of undamaged muscle tissue. A necrotic lesion is visible in the histology section as an area of fragmented and damaged fibers with infiltration by purple-stained inflammatory cells (Fig. 5(e)). As in Fig. 4, in the OCT image this corresponds to an area which lacks the striation pattern of undamaged muscle fibers (Fig. 5(g)) [37], and in the micro-elastogram, this same area not only lacks a striation pattern, but additionally possesses a mottled texture (Fig. 5(i)). Once again, the contrast between intact fibers and necrosis is stronger in the micro-elastograms than in the OCT images, demonstrating OCME’s greater ability to distinguish between necrotic and intact muscle tissue.
Fig. 5 Images of the left gastrocnemius muscle from an unexercised mdx mouse. En face (a) H&E-stained histology section, (b) OCT image and (c) micro-elastogram. (d), (f) and (h) Magnified insets (2 × 2 mm) of corresponding regions of intact muscle fibers. (e), (g) and (i) magnified insets of corresponding regions of a necrotic lesion. OCT images and micro-elastograms were taken 100 µm from the surface.
Figure 6 shows micro-elastograms of an excised left gluteus muscle from the same mdx mouse used to generate the images in Fig. 5. This figure further demonstrates the capability of OCME to distinguish necrotic lesions from areas of intact muscle tissue. Visible in this figure are areas of undamaged muscle fibers in the H&E-stained histology (Fig. 6(d)), corresponding to areas with longitudinal striation patterns in the OCT image (Fig. 6(f)) and micro-elastogram (Fig. 6(h)). Also visible is a necrotic lesion, seen as an area with purple inflammatory cells in the histology (Fig. 6(e)), as an area without striations in the OCT image (Fig. 6(g)), and as an area with a mottled texture in the micro-elastogram (Fig. 6(i)).
Fig. 6 Images of the right gluteus muscle from the same unexercised mdx mouse as Fig. 5. En face (a) H&E-stained histology section, (b) OCT image and (c) micro-elastogram. (d), (f) and (h) magnified insets (2 × 2 mm) of corresponding regions of intact muscle fibers. (e), (g) and (i) Magnified insets of corresponding regions of a necrotic lesion. OCT images and micro-elastograms were taken 200 µm from the surface.
3.4. Micro-scale strain contrast in skeletal muscle
To further examine the contrast provided by OCME, Fig. 7 presents plots of OCT SNR (Figs. 7(g), 7(h) and 7(i)) and local axial strain (Figs. 7(j), 7(k) and 7(l)) obtained from scans of gastrocnemius and gluteus muscles of two C57 wild-type mice. These plots, taken from a depth of 100 µm, show the variation in OCT SNR and strain across a lateral range of 2 mm, and were averaged in the orthogonal lateral direction over 160 µm (red-dashed regions in Figs. 7(a)–7(f)). The spacing between the small dips in strain (Figs. 7(j), 7(k) and 7(l)) is visible as longitudinal striations in the en face images (Figs. 7(b), 7(d) and 7(f)), and shown by the red markings in Figs. 7(g)–7(l), is on the order of 50 µm. This is consistent with the size and architecture of individual muscle fibers, which range from approximately 20–100 µm in diameter [1, 35], and are arranged in parallel. The spacing between the large dips in strain, visible as the strong striation patterns in the en face images, and shown by the blue markings in Fig. 7, is on the order of 200–500 µm. This is consistent with the size of muscle fascicles, which comprise several muscle fibers enclosed by a sheath of connective tissue (perimysium) [1]. The large compressive strain measured at the boundary of the fascicles suggests that the perimysium is much softer under compressive loading than muscle fibers. This is discussed further in Section 4. The OCT images show longitudinal striations which are consistent with the size of muscle fibers, or small groups of muscle fibers, but do not show clearly the large striation patterns expected from the muscle fascicles.
Fig. 7 Averaged plots of OCT SNR and local axial strain from three C57 muscles: (a), (b), (g) and (j) right gastrocnemius; (c), (d), (h) and (k) left gastrocnemius; (e), (f), (i) and (l) left gluteus, from the same mouse as the left gastrocnemius. En face 2 × 2 mm regions: (a), (c) and (e) OCT images and (b), (d) and (f) micro-elastograms. (g), (h) and (i) averaged OCT SNR over the regions marked in (a), (c) and (e), respectively. (j), (k) and (l) averaged local axial strain over the regions marked in (b), (d) and (f), respectively. Blue markings indicate representative widths of the striation patterns formed between lines of strong compressive strain. Red markings indicate representative widths of the striation patterns formed between lines of less intense compressive strain. OCT images and micro-elastograms were taken 100 µm from the surface.
To evaluate the contrast provided by OCT and OCME, we define the striation contrast, Sc, as the ratio between the signal at the peak of the striation, sp, to the signal at the trough of the striation, st, expressed on a log scale as Sc = |20log10(sp/st)|, following the convention used in previous studies of OCE [41] and ultrasound elastography [47]. In addition, for OCME, only points with negative (compressive) strain are considered. The striation contrast is evaluated for the averaged plots from the right gastrocnemius sample shown in Figs. 7(g) and 7(j), the left gastrocnemius sample shown in Figs. 7(h) and 7(k), and the left gluteus sample shown in Figs. 7(i) and 7(l). The striation contrast for the OCT plots is 4.5–6.2 dB, 8.0–9.1 dB, and 5.0–7.5 dB (95% confidence interval of the mean), respectively. The contrast for the large striations in the micro-elastogram plots is 22–34 dB, 12–17 dB, and 19–22 dB, respectively. Similarly, the contrast for the small micro-elastogram striations is 6.6–9.0 dB, 3.1–7.5 dB, and 7.0–10 dB, respectively. These results demonstrate that OCME can provide high contrast images of muscle structure, which are complementary to OCT.
4. Discussion
The results presented in this study demonstrate the ability of OCME to provide micro-scale mechanical contrast of skeletal muscle tissue. Skeletal muscle consists of a hierarchy of filamentous structures [1]. Individual muscle fibers (myofibers) are surrounded by a sheath of connective tissue, the endomysium, and bound to adjacent fibers to form fascicles, which are enclosed within another, thicker, sheath of connective tissue, the perimysium. Previous studies have shown that the muscle extracellular matrix, which includes the endomysium and perimysium, is much stiffer than the muscle fibers when subjected to tensile forces [48]. However, both sheaths are primarily made of collagen fibers, which are strong under tension, but relatively soft under compression [1, 49]. The results of this study suggest that, under compression, the sheaths appear softer than the muscle fibers, leading to the striation patterns observed in the micro-elastograms. The results shown in Section 3.4 demonstrate that the striation patterns in the micro-elastograms can provide equal or greater contrast (~12–34 dB for the large striations, ~3–10 dB for the smaller striations) than can the OCT images (~4–9 dB). However as seen in Fig. 7, the patterns in the micro-elastograms do not exactly correspond to the patterns in the OCT images. The striation patterns observed in the OCT images are consistent with scattering due to changes in refractive index between the muscle fibers and the connective tissue sheaths. This may explain why fascicles are not apparent in the OCT images, as the myofiber-endomysium boundary may not be sufficiently optically distinct from the myofiber-perimysium boundary. This distinction is much stronger in micro-elastograms, since the greater thickness of the perimysium allows it to be more readily distinguished from the muscle fibers than the thinner endomysium, leading to the large dips in strain seen in this study (Fig. 7). Mechanical interaction within the interfascicular space, for example, muscle fibers sliding past one another under compressive loading, could also be a cause of the large compressive strain observed between fascicles in this study.
Figures 4, 5, and 6 demonstrate the capability in micro-elastograms to identify regions of necrosis in skeletal muscle samples from mdx dystrophic mice. The necrotic lesions appear distinct from the surrounding, intact, muscle tissue, and this contrast is likely due to the difference in structure and mechanical properties of the necrotic lesions compared to the intact muscle tissue; the pathology forms regions of broken fibers and inflammatory cells [35]. Necrotic lesions often possess a slightly mottled appearance in the micro-elastograms (for example, in Fig. 4), which is likely due to the inherent mechanical and structural heterogeneity of necrotic lesions, as seen in the H&E-stained histology (Figs. 4(d) and 4(e)). From the current results, it is unclear whether this mottled appearance is correlated with the severity of necrosis, but this could be an avenue for future studies.
Muscle tissue is highly mechanically anisotropic; thus, the sample orientation is important in determining the quality of the resulting micro-elastograms. We observed that the best results were obtained when the muscle fibers were oriented longitudinally to the en face plane and, thus, we ensured that all samples were oriented in roughly this manner prior to imaging. The architecture of skeletal muscles is complex, particularly in muscles such as the triceps and quadriceps, which comprise multiple large muscle bundles arranged at distinct angles relative to each other [1]. This can be seen in the upper-right portion of Fig. 3, where the fibers shift from being oriented parallel to the en face plane to being oriented almost perpendicular to the plane. In this perpendicular orientation, the strain contrast between cross-sections of fibers and fascicles produces a pattern of interlaced focal spots of high and low strain in the micro-elastograms, distinct from the striation patterns shown in the insets in Figs. 2–7. In this study, we have carefully positioned the samples to minimize this effect; however, further study is required to judge the full extent to which sample orientation affects the ability to distinguish necrotic lesions from intact muscle tissue.
An inherent assumption of OCME is that the sample compresses in response to uniaxial loading. Based on our method of processing [25, 41], the resulting local strain should ideally be negative, indicating compression. In some instances, however, positive strain is measured. This effect, also noted in our previous study on breast tissue [25], tends to occur mostly at the edges between distinct features in the en face plane, for example, between muscle fibers, or at the boundary of a duct in the case of breast tissue. Thus, positive strain can help to accentuate the contrast visible in micro-elastograms, but could complicate attempts to quantify elasticity. We hypothesize that one cause of this positive strain is the combined heterogeneity and incompressibility of tissue; i.e., compression of one region of tissue results in the expansion (and, therefore, positive strain) of an adjacent region of tissue to conserve volume. We have observed that reducing the compressive preload force applied before imaging can help reduce this effect. Further study is required to investigate this phenomenon, and we are currently implementing finite element models of sample deformation to elucidate this aspect of OCME [41].
The mechanical contrast shown in this study was generated using an external compressive load, creating micro-elastograms based on the passive mechanical properties of muscle tissue. Muscle additionally possesses active mechanical properties, which are typically measured on a macro-scale through stimulus of the muscle and functional testing of the resulting forces [3, 50, 51]. In principle, it should be possible to generate elastographic contrast from active mechanical function of the muscle, for example, by measuring the lateral strain generated in response to an electrical stimulus. In conjunction with a faster acquisition method [52], this would allow micro-elastograms to be directly correlated with functional measurements of muscle mechanical properties, which could be highly useful in physiological studies of skeletal muscle pathology. In addition, a variant of OCE which we have recently developed, optical palpation [53], could be used to provide high-resolution measurements of the forces generated by muscles under active contraction. Whilst this current study has been performed on excised tissue samples, it is feasible that such techniques may be extended to allow in situ, or possibly in vivo, measurements of muscle mechanical properties. For in vivo measurements to be feasible, the loading mechanism needs to be able to penetrate deeply enough to reach muscles which are possibly occluded by other tissues and, if penetrating tissue, of sufficiently small cross section to minimize harm to the subject. This could be achieved with needle-based OCE [23, 54], but further study is required.
The micro-elastograms presented in this study are qualitative maps reflecting the relative mechanical contrast provided by strain imaging, rather than quantitative measurements of tissue elastic modulus. Related OCE techniques, such as those based on surface acoustic [19] or shear waves [26, 34], provide, within experimental error, absolute measurements of elastic modulus; however, these techniques have yet to demonstrate the ability to resolve the fine structures visible in the micro-elastograms presented in this study [17, 25]. To provide quantitative elasticity values using OCME requires knowledge of both the stress and strain distribution throughout the sample. A step towards this goal may be possible by combining OCME with optical palpation [53]. This combination could yield, at high-resolution, the stress distribution across the surface of a tissue sample. It may then be possible to deduce the stress distribution throughout the tissue by combining OCME with optical palpation and applying some form of inverse method.
5. Conclusion
In conclusion, we have shown that OCME can provide micro-scale images demonstrating the mechanical contrast in skeletal muscle tissue from the mdx mutant mouse model of DMD and from wild-type C57 mice. Through comparison with OCT and co-registered histology, we have shown that OCME can distinguish tendons from muscles and necrotic lesions from intact muscle tissue. We observe that the longitudinal striation patterns visible in micro-elastograms are consistent with the size and arrangement of clusters of muscle fibers (fascicles), as well as individual muscle fibers. Many of these striations are not visible in the OCT images, demonstrating the additional contrast provided by OCME. By probing tissue mechanical properties on the intermediate length scale, between that of cells and whole organs, OCME provides opportunities to enhance the understanding of the interaction between such properties and the development and progression of pathologies.
Acknowledgments
The authors thank CELLCentral, The University of Western Australia, for preparation of many of the histology slides, and Jeremy Costin, for helping with matching of OCT images and micro-elastograms to histology. The authors acknowledge the facilities, and the scientific and technical assistance of the Australian Microscopy & Microanalysis Research Facility at the Centre for Microscopy, Characterisation and Analysis, The University of Western Australia, a facility funded by the University, State and Commonwealth Governments. L. Chin is supported by a Robert and Maude Gledden postgraduate research scholarship from The University of Western Australia. K. M. Kennedy is supported by a Scholarship for International Research Fees, The University of Western Australia. P. Wijesinghe is supported by a William and Marlene Schrader Scholarship, The University of Western Australia. This project is supported with funding from the Australian Research Council; Cancer Council WA; National Breast Cancer Foundation, Australia; and the National Health and Medical Research Council, Australia.
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