Collagen fibril abnormalities in human and mice abdominal aortic aneurysm

Abstract

Vascular diseases like abdominal aortic aneurysms (AAA) are characterized by a drastic remodeling of the vessel wall, accompanied with changes in the elastin and collagen content. At the macromolecular level, the elastin fibers in AAA have been reported to undergo significant structural alterations. While the undulations (waviness) of the collagen fibers is also reduced in AAA, very little is understood about changes in the collagen fibril at the sub-fiber level in AAA as well as in other vascular pathologies. In this study we investigated structural changes in collagen fibrils in human AAA tissue extracted at the time of vascular surgery and in aorta extracted from angiotensin II (AngII) infused ApoE^−/− mouse model of AAA. Collagen fibril structure was examined using transmission electron microscopy and atomic force microscopy. Images were analyzed to ascertain length and depth of D-periodicity, fibril diameter and fibril curvature. Abnormal collagen fibrils with compromised D-periodic banding were observed in the excised human tissue and in remodeled regions of AAA in AngII infused mice. These abnormal fibrils were characterized by statistically significant reduction in depths of D-periods and an increased curvature of collagen fibrils. These features were more pronounced in human AAA as compared to murine samples. Thoracic aorta from Ang II-infused mice, abdominal aorta from saline-infused mice, and abdominal aorta from non-AAA human controls did not contain abnormal collagen fibrils. The structural alterations in abnormal collagen fibrils appear similar to those reported for collagen fibrils subjected to mechanical overload or chronic inflammation in other tissues. Detection of abnormal collagen could be utilized to better understand the functional properties of the underlying extracellular matrix in vascular as well as other pathologies.

GRAPHICAL ABSTRACT

1. INTRODUCTION

The extracellular matrix (ECM) plays a major role in defining the architecture of the vessel wall and its underlying functional properties. Collagen and elastin are the major components of the vascular ECM and organize the vessel wall into its three distinct layers (intima, media and adventitia), along with the resident endothelial cells, smooth muscle cells (SMC) and adventitial fibroblasts respectively[1]. Abdominal aortic aneurysms (AAA)[2] is a vascular disease characterized by dilatation of the abdominal aorta accompanied by extensive remodeling of the ECM [3,4]. Elastin degradation, along with an imbalance in collagen synthesis and degradation[5,6] is a well-established feature of ECM remodeling in AAA [4,7,8]. However besides their quantity, it is the macromolecular organization and structure of elastin and collagen fibers/fibrils that dictate the functional characteristics of the aneurysm such as its stiffness[9], susceptibility to rupture[10], extent of mineralization[11], and thrombus formation[12,13].

A number of studies by us[14][15] and others[16] [17][18] have elucidated disruptions of the elastic lamina in AAA by using histological staining in murine models as well as in clinical AAA. Ultrastructural investigations have provided further insights into macromolecular structural changes. For instance transmission electron microscopy (TEM) analysis has revealed the presence of nascent elastin fibrils lacking fibrilin-1, which results in deposits of amorphous elastin but not mature elastic fibers in an elastase-infused mouse model of AAA [17]. In clinical AAA elastin degradation is known to result in fragmented elastin fibers[16] and electron-translucent elastin, (often only detectable by immunogold labeling or electron backscatter analysis)[19]. Besides AAA, similarly disrupted elastic fibers are also found in other vascular diseases like thoracic aortic aneurysm and Marfan’s syndrome[19,20]. It is interesting to note that an aberrant elastin remodeling have also been observed in the dermal tissue of patients with certain vascular diseases like spontaneous cervical artery dissection (sCAD), internal carotid dissection, and vertebral artery dissection[21,22].

The organization of collagen at the fiber level is also understood to be affected in AAA. Scanning electron microscopy and polarized light microscopy have revealed that adventitial collagen fibers in AAA are less undulating (wavy) as compared to control samples[23]. A similar observation has been reported using second-harmonic generation (a multiphoton imaging based technique) analysis, where the adventitial collagen fibers exhibited reduced waviness or crimp in clinical AAA[24] and in a mouse model of AAA[25]. However, very little is understood about the changes at the sub-fiber level i.e. in the fibril structure of collagen in AAA and other vascular diseases. This is especially important as the collagen fibril is the precursor to macromolecular ECM remodeling and the fibril structure dictates not only the fiber properties but also cell-matrix interactions and functional characteristics of the aneurysm as stated above.

Our current knowledge regarding alterations in the collagen fibril structure in vascular diseases has been primarily derived from studies on dermal connective tissue of these patients. For instance, reduced fibril diameters have been reported in dermal tissues from patients with vascular Ehlers-Danlos Syndrome (EDS type IV) and intracranial aneurysms[26]. Another feature, namely irregular cross-sectional profiles (which depart from the normally circular cross-section) of collagen fibrils have been found in the dermal tissues from a variety of vascular diseases like EDS IV[27], sCAD, internal carotid, vertebral artery dissection[21,22,28] and intracranial aneurysms[26]. And finally collagen fibrils with diminished or nearly-absent D-periodic banding are also present in the skin of EDS type IV[29] and sCAD patients[30].

The evidence of collagen fibril abnormalities occurring in dermal tissues of patients with vascular diseases prompted us to examine if collagen fibril abnormalities are also present in the pathological vascular tissues. This is especially pertinent in AAA as several factors including, mechanical overload, inflammation, enzymatic digestion by collagenases as well as factors which can perturb collagen fibrillogenesis are known to be present in AAA. To determine ultrastructural differences in collagen fibril structure, we used clinical AAA tissue obtained during vascular surgery as well as a well-established mouse model of AAA resulting due to angiotensin II (AngII) infusion[8,31,32]. Control non-AAA samples from each species was utilized for comparison. Two independent ultrastructural microscopy techniques, namely, transmission electron microscopy (TEM) and atomic force microscopy (AFM), were used to characterize collagen fibril morphology. Image analysis was performed to quantify parameters like length and depth of D-periods, fibril diameter and curvature. We elucidate how collagen fibril abnormalities are present in AAA, which can be potentially utilized to understand the pathogenesis of AAA and functional role of the underlying ECM.

2. MATERIALS AND METHODS

2.1. Mice, Aneurysm Model, Experimental Groups.

All the animal-related experiments were conducted via protocols (#8799 and #AR11–00031) approved by the Animal Care and Use Committee (ACUC) at the University of Missouri (Columbia, MO) and the Institutional Animal Care and Use Committee (IACUC) of the Research Institute at Nationwide Children’s Hospital (Columbus, OH) respectively. All the animal experiments conformed to the NIH guidelines (Guide for the Care and Use of Laboratory Animals). An AngII-infused mouse model of AAA was used for this study [8,31,32]. Only male mice were studied for the in vivo studies because of low incidence of AngII-induced AAA in female mice[33]. Briefly, at 8–10 weeks of age male ApoE^−/− knockout mice (C57BL/6J background (Jackson Laboratory, Bar Harbor, ME) were infused with AngII (1000 ng/kg/min) or saline from a subcutaneously implanted mini-osmotic pump (model 2004; Alzet, Cupertino, CA). For implantation of mini-osmotic pumps, mice were anesthetized in a closed chamber with 3% isoflurane in oxygen for 2 to 5 minutes until immobile as previously described[31]. Each mouse was then removed, and taped on a heated (35–37°C) procedure board with 1.0–1.5% isoflurane administered via nosecone during minor surgery. After 28 days of AngII (or saline) infusion, the mice were euthanized with an overdose of ketamine/xylazine (100 and 20 mg/kg, respectively) and the entire aorta was extracted. Thereafter segments of the suprarenal abdominal aorta and distal portion of the thoracic aorta were utilized for analysis. AAA was confirmed as a dilated segment of the suprarenal aorta and by histological evaluation. A total of n= 8 AngII infused mice and n=6 shams (saline infused mice) were utilized for collagen fibril analysis as detailed below.

2.2. Clinical Tissue.

Human AAA tissue was obtained in accordance with an approved Institutional Review Board protocol (#2015H0233 approved 10/26/2015) from The Ohio State University. All investigations conformed to the principles outlined in the Declaration of Helsinki. The patients were recruited at the Ross Heart Hospital during the period January 2017 to February 2020. At this site, approximately 300 patients with infrarenal AAA are examined annually and approximately 50 undergo either minimally invasive or open surgery every year. Patients who were diagnosed with AAA over 5.5 cm in diameter (measured on CT scan) and who were judged to be candidates for elective open aneurysm repair by the treating surgeon were considered eligible. Surgery was recommended in accordance with the Society for Vascular Surgery guidelines recommending repair of AAA when the diameter reaches 5.5 cm in good risk patients. Clinical tissue was obtained after a written informed consent from patients prior to undergoing surgery to treat AAA per standard of care. All eligible patients were asked to participate in the study without any bias and those who consented were recruited for this study. Tissue for this study was obtained from n=6 male patients, in the age group 65 to 75 years and they self-identified themselves as belonging to the non-Hispanic white population. After AAA repair, a ~3 × 3 cm segment of anterior aortic sac was excised and utilized for the following studies.

Non-aneurysmal control samples were collected from the infrarenal segment of aorta (n=6) at autopsies within 24 hours of death at the Detroit Coroner’s Office as described in our previous study[34]. The samples were considered discarded tissues and no consent was required. At the time of harvesting samples, no link to personal identifying information was kept. The collection of the tissue samples and their use for research projects was approved by the Institutional Review Board of Wayne State University, Detroit, Michigan, and the research carried out was in compliance with the Helsinki Declaration. The number of samples (study size) was determined by a prospective power analysis performed using α=0.05 and power=0.95. Based on our earlier studies on analysis of depth of D-periods in collagen fibrils in the murine aorta[35], we estimated a standard deviation of 0.3 nm and sought to detect a difference of at least 1 nm between the D-period depth of normal and abnormal collagen in clinical human AAA tissue. Based on these inputs, our power analysis indicated a sample size of n=5 for clinical AAA group.

2.3. Histology.

Segments of the murine and human aortic tissue samples were incubated in phosphate-buffered formalin and embedded in paraffin for histological analyses. Serial sections (5 μm) of the abdominal aorta were subjected to Verhoeff–Van Gieson stain (VVG; elastin) and picrosirius red stain (collagen) for histoarchitectural evaluation of aneurysm as described previously[14][15]. Birefringence from collagen fibers in picosirius red stained samples was analyzed using a polarized light microscope (Leica DM5500) to evaluate the content of thick (red-orange) or thin (green) fibers[36].

2.3. Transmission electron microscopy (TEM).

For TEM analysis, segments of suprarenal abdominal aorta or thoracic aorta from mice (n=6 AngII; 6 saline-infused) and clinical AAA or control human tissue (n=6 AAA; 6 control) were dissected, fixed in 4% glutaraldehyde for at least 24 hours, and processed for routine TEM as previously described[35]. Briefly, aortic pieces were osmium tetroxide post-fixed, dehydrated in a graded series of ethanol, and embedded in Spurr’s epoxy resin (Electron Microscopy Sciences). Thin sections (~ 94 nm in thickness)) were cut and imaged using a JOEL JEM-1400 TEM equipped with a SIS Olympus MegaView III digital camera (with pixel size of its charged coupled device (CCD) detector as 6.45 × 6.45 μm). Images were acquired at 1376 × 1032 pixels with magnifications ranging from 3k to 150k. Multiple regions spanning the intima to the adventitia were imaged in murine aortic sections to assess ECM remodeling, collagen morphology and location of abnormal collagen. For analysis of abnormal collagen, images were acquired from at least five different areas in the remodeled regions in both murine and human samples. TEM imaging and identification of abnormal collagen (with weakened D-periodicty) was performed by three independent users.

2.4. Atomic force microscopy (AFM).

Vascular tissue pieces adjacent to that used for TEM were fixed in 2% glutaraldehyde, embedded in Optimal Cutting Temperature (OCT) compound, flash frozen in liquid nitrogen, and stored at −80°C until further use. For AFM studies, the OCT embedded tissue was cryosectioned into 5 μm thick sections and mounted onto poly-lysine coated glass cover slips. After washing with ultrapure water and air drying, sections were imaged using a Multimode AFM equipped with a JV scanner and a Nanoscope iIIa Controller (Digital Instruments, Santa Barbara). Images (with 512 pixels per scan direction) were acquired in tapping mode in ambient air using NSC15 cantilevers (Mikromasch) as previously described and calibrated[35]. Adventitial collagen and regions showing ECM remodeling could be easily identified in the tissue sections using a reflected light module coupled to the AFM equipment. AFM height and amplitude images were acquired from these remodeled regions.

2.5. Image analysis.

Fibril diameters were determined using ImageJ (NIH) using longitudinal and cross sectional images of collagen fibrils as stated. To determine the percent abnormal collagen, ImageJ (NIH) was used to trace total collagen area and total abnormal collagen area from TEM images of remodeled adventitia. This fraction determined for each image (n=5 per sample) was used to determine percent abnormal collagen in each sample. Fibril curvature was determined using FiberApp, a Matlab-based software. FiberApp is based on statistical polymer physics and enables analysis of fiber-like, filamentous, and macromolecular objects[37]. Individual collagen fibrils in TEM images were manually traced and the local curvature (1/μm) of the fibrils along their contour length was ascertained using a step size of 3.7 nm. The length and depth of D-periods in collagen fibers was ascertained using section profile analysis on height images using the Nanoscope software as previously described [35].

2.6. Statistical Analysis.

Data are presented as means ± standard deviations. Statistically significant differences in fibril diameters, fibril curvature, and length and depth of D-periods in normal vs. abnormal collagen fibrils from AngII- vs. saline-infused mice (4 group comparison: normal fibrils in abdominal aorta of saline-infused; normal fibrils in thoracic aorta from AngII-infused; normal fibrils in abdominal aorta of AngII-infused and abnormal fibrils in abdominal aorta of AngII-infused mice), one way ANOVA was utilized and when necessary followed by pairwise comparison of means by Tukey-Kramer test. The data for each of these tests complied with the constraints of the test as validated by normality and equal variance tests. To ascertain statistically significant differences for the same variables in normal vs. abnormal collagen fibrils from clinical human AAA vs. control samples (3 group comparison: normal fibrils in control and AAA samples and abnormal fibrils in AAA samples), one way ANOVA was again utilized and when necessary followed by pairwise comparison of means by Tukey-Kramer test. The data for each of these tests complied with the constraints of the test as validated by normality and equal variance tests. Due to inter-sample variability, a matched pairs t-test was also used as a secondary test to compare normal vs. abnormal collagen within murine or human AAA. The data for each of these tests was found to be approximately normally distributed and free of outliers. All statistical analysis was performed using SAS JMP software (version 14.0), and a p- value <0.05 was considered significant.

3. RESULTS

AngII-infused mice exhibited abnormal dilation of the suprarenal aorta characteristic of AAA (Figure 1, A, inset). Human AAA was clinically established and crude tissue was extracted at the time of surgery (Figure 1, B, inset). Verhoeff-van Gieson stain and TEM analysis confirmed the presence of intact elastic lamina and adventitial layers in saline-infused mice (Figure 1, A and Figure S1, A-C). Aneurysms in AngII-infused mice exhibited diminished and disrupted elastic lamina and a remodeled adventitia (Figure 1, A and Figure S1, D-E) as previously reported [8,31]. Similar observations were made for human control and AAA samples (Figure 1, B). Picrosirius red staining of aortic sections showed increased collagen content in AngII-infused mice and in human AAA tissues (Figure 1, C and D) as compared to their respective control samples. Interestingly, the remodeled regions in AngII-infused mice and clinical AAA were abundant in both thick (red-orange) and thin (green) collagen fibers

Figure 1: Histological staining of aortic tissue to evaluate elastin and collagen content.

Insets (center of panel A) shows dilatation of suprarenal aorta in AngII-infused mice and (center of panel B) a ~3 × 3 cm patch of crude AAA tissue harvested from a patient undergoing vascular surgery. Abdominal aortas from saline or AngII-infused ApoE^−/− mice (A) and from human (non-AAA) donor controls and clinical AAA (B) were stained with Verhoeff-van Gieson. Elastin is stained brown and both AngII-infused mice and human AAA exhibit depletion and fragmentation of elastin fibers in remodeled regions. Similar samples (as in A and B) were stained with picrosirius red (C and D) and examined using polarized light. Thick collagen fibers appear as red-orange while thin fibers appears green. Both AngII-infused mice and human AAA display significant remodeling and increased collagen content. . All scale bars are 50 mm.

3.1. Abnormal Collagen Fibrils.

To examine the ultrastructural morphology of collagen fibrils, we examined the D-periodic banding in longitudinal sections of collagen fibrils in TEM images. The D-periodic banding in collagen fibrils is characteristic of its native molecular packing and arises due to the gap and overlap regions resulting from staggered arrangements of collagen molecules[38]. ‘Normal’ collagen fibrils with well-defined characteristic D-periodicity, nearly straight contours and parallel alignment within fibril bundles could be easily identified in the adventitia of abdominal aorta from both saline and AngII-infused mice (Figure 2, A, B, D, E and G, H). A similar feature of normal collagen fibrils was observed in the adventitia of distal portion of thoracic aorta (TA) from AngII-infused mice (Figure S2, A–C). However the abdominal aorta from AngII infused mice also contained several pockets of ‘abnormal’ fibrils interspersed between bundles of normal fibrils (Figure 2, C, F and I). These abnormal fibrils exhibited D-periods with a reduced contrast, or a complete lack of it, along with a “wavy” or undulating appearance. Such abnormal collagen fibrils were primarily found in the heavily remodeled regions consisting of disrupted external elastic lamina and adventitia (Figure 3) in the abdominal aorta of AngII-induced mice and were not observed in the abdominal aorta from saline-infused mice or distal portion of the thoracic aorta from Ang-II induced mice, both of which were devoid of visible adventitial thickening.

Figure 2: Abnormal collagen fibrils in AngII-infused mice.

TEM images of adventitial collagen in murine aortic sections from Ang-II or saline infused mice as indicated. Collagen fibrils with normal D-periodic banded structure were observed in both treatment groups (A-B, D-E, G-H). However, Ang-II infused mice also revealed a subset of abnormal fibrils with disrupted or unresolvable D-periods (C, F, I). Low magnification images show D-periodic banded collagen fibrils with straight contours (A-B), while abnormal collagen fibrils showed undulating or wavy contours (C). All scale bars are 200 nm. TEM images of abnormal collagen fibrils from additional AngII-infused mice are shown in Figure S3.

Figure 3: Location of abnormal collagen fibrils in AngII-infused mice.

TEM images show area of disrupted external elastic lamina where adventitial and medial collagen are no longer clearly segregated and bundles of normal adventitial collagen are present on both medial and adventitial sides (blue arrows in A). Interspersed amongst this normal collagen, pockets of ‘abnormal’ collagen with a diminished D-periodicity are also present (red arrows in A). Higher magnification images of selected areas consisting of normal (blue arrows) and abnormal (red arrows) collagen are shown in (B-C) and (D-E) respectively. All scale bars are 200 nm.

Similar features were identified in human control and clinical AAA, wherein like the murine tissues normal collagen fibrils with straight contours and well-defined characteristic D-periodicity could be identified (Figure 4, A, B, D, E and G, H). The clinical AAA tissue also contained several regions of abnormal fibrils exhibiting a “wavy” or undulating appearance, with a reduced contrast or a complete lack of D-periodicity (Figure 4, C, F and I) even more severe than that found in the AngII-infused mice. TEM images from additional samples showing abnormal fibrils with compromised D-banding in clinical AAA and AngII-infused mice are shown in Figure S3. The abnormal collagen fibrils could be easily found in the remodeled ECM of all AAA samples examined. Analysis of n=5 TEM images per sample revealed that the fraction of abnormal collagen to total collagen present in a remodeled region ranged from ~15 to 55% in human and ~10 to 45% in murine AAA tissues.

Figure 4: Abnormal collagen fibrils in human AAA.

TEM images of adventitial collagen fibrils in clinical AAA tissue as well as healthy controls show fibrils with normal D-periodic banded structure (A-B, D-E, G-H). However, clinical AAA tissues also displayed abnormal fibrils with disrupted or unresolvable D-periods (C, F, I). Low magnification images show D-periodic banded collagen fibrils with straight contours (A-B), while abnormal collagen fibrils showed undulating or wavy contours (C), similar to those observed in murine AAA. All scale bars are 200 nm. TEM images of abnormal collagen fibrils from additional clinical AAA samples are shown in Figure S3.

3.2. Fibril diameter.

We next evaluated the diameters of adventitial collagen fibrils in AngII-and saline-infused mice, and in human controls and clinical AAA, using longitudinal sections of fibrils in TEM images. Before analyzing collagen in AAA, baseline comparisons of medial and adventitial collagen appearance and diameter were performed on saline-infused (control) murine abdominal aortas. As shown in Figure S4, the medial collagen (~ 32 nm) was nearly half the fibril diameter as compared to adventitial collagen (~ 53 nm) and exhibited reduced contrast in D-periodicity and an undulating appearance similar to that exhibited by abnormal collagen fibrils shown in Figure 2. However, in the remodeled regions of AngII-infused mice, both normal and abnormal fibrils had an average diameter of ~ 53 nm (Figure 5, A and B) with no statistically significant differences between their mean values (one way ANOVA, p=0.4872). Human adventitial collagen in non-AAA controls samples, had an average fibril diameter of ~47 nm, slightly lower than that of murine collagen. As observed in murine samples, when comparing abnormal fibrils (in AAA) to normal fibrils (in control and AAA) there was no statistically significant differences in their mean values (Figure 5, C and D, one way ANOVA, p=0.2796). However, it should be noted that in AAA samples, the abnormal fibrils exhibited a small (~ 15%) but statistically significant reduction in their average fibril diameter when using a matched pairs t-test for human samples (p=0.0324) but not murine samples (p=0.4177). Thus the change in fibril diameter between normal and abnormal fibrils was minimal and both types of fibrils resembled adventitial collagen in their diameters.

Figure 5: Collagen fibril diameter in AngII-infused mice and human AAA.

Distribution of fibril diameters from a representative murine and human sample as indicated (A, C). Longitudinal sections of adventitial collagen fibrils in TEM images of abdominal aorta were utilized for diameter analysis. Average fibril diameters from each sample were compared for normal and abnormal fibrils for all murine (B) and human samples (D) as indicated. No significant difference was found in collagen fibril diameter between the normal fibrils in saline or AngII-infused mice or between normal and abnormal fibrils in AngII-infused mice (p=0.4872) (B). An ANOVA for the human sample groups also indicated no significant difference in collagen fibril diameter (p=0.2796); however, pairwise comparison of human AAA samples indicated that the abnormal collagen fibrils in AAA had a small but statistically significant reduction in fibril diameter as compared to normal fibrils in the corresponding samples (p=0.0324) (D).

3.3. Fibril Curvature.

As previously noted, abnormal collagen fibrils in both AngII infused mice and human AAA clinical samples, exhibited undulating contours compared to the straight contours exhibited by normal collagen fibrils. We therefore quantified and compared the local fibril curvature of collagen fibrils using image analysis (Figure 6). In mice, normal collagen fibrils in both the AngII and saline models exhibited fibrils with an average curvature of 0.9 ± 0.2 μm⁻¹ and 1.0 ± 0.3 μm⁻¹ respectively, while abnormal collagen fibrils in the AngII-infused mice exhibited fibrils with a significantly higher degree of curvature averaging at 3.1 ± 0.5 μm⁻¹ (Figure 6, A and B one way ANOVA followed by Tukey-Kramer test, p˂0.0001). Similarly, human AAA exhibited abnormal collagen fibrils with a significantly higher degree of curvature than their normal fibril counterparts, averaging 4.4 ± 0.7 μm⁻¹ as compared to 1.4 ± 0.1 μm⁻¹ and 2.1 ± 0.5 μm⁻¹ for normal collagen in human controls and AAA respectively (Figure 6, C and D, one way ANOVA followed by Tukey-Kramer test, p˂0.0001). We thus elucidate that abnormal collagen fibrils in AAA were characterized by an increased curvature as compared to normal fibrils.

Figure 6: Collagen fibril curvature in AngII-infused mice and human AAA.

Normal and abnormal adventitial collagen fibrils in TEM images were traced and fibril curvature was determined for individual fibrils. Histograms showing distribution of collagen fibril curvature from a representative murine (A) and a human (C) sample as indicated. Abnormal collagen fibrils found in AngII-infused mice aortas exhibited significantly (*p˂0.0001) greater average curvature than normal fibrils found in either saline-or AngII-infused mice (B). Similarly, in human AAA abnormal collagen fibrils had a significantly (*p˂0.0001) greater average curvature than normal fibrils in either AAA or controls (D).

3.4. D-depth.

To derive a quantitative parameter for lack of D-periodicity, we imaged collagen fibrils using AFM, a high-resolution technique which can track nanoscale topography of samples. Consistent with our TEM observations, AFM amplitude images revealed well-defined D-periodic bands in normal collagen fibrils present in the adventitia of both saline and AngII-infused mice (Figure 7, A and B) as well as in human controls and AAA (Figure 7, D and E). However, similar to TEM observations, we also observed abnormal collagen fibrils with compromised D-periodic contrast in AFM images of AngII-infused mice (Figure 7, C) and in human AAA (Figure 7, F). Additional examples of abnormal collagen fibrils in AngII-infused mice and human AAA can be found in Figure S5. No differences in the length of D-periods in (66 ± 6 nm, one way ANOVA, p=0.473) were observed between the normal collagen fibrils present in AngII or saline infused mice or in human AAA samples. The length of D-periods in abnormal fibrils (when measureable) also resembled that of normal collagen fibrils. To quantify the reduced contrast of D-periodicity, we also measured the depth of D-periods of collagen fibrils from AFM height images (Figure 8 and 9). There was no difference in the mean D-depth of normal collagen fibrils between AngII-and saline-infused mice (Figure 8, A–D, Tukey-Kramer test, p=0.973). However, a significant decrease in D-depth was observed in the abnormal collagen fibrils in AngII-infused mice (Figure 8, E and F, one way ANOVA followed by Tukey-Kramer test, p˂0.0001), and in certain cases the depth could not be measured. Similarly, in human AAA, a significant decrease in D-depth was found only for abnormal collagen fibrils (Figure 9, one way ANOVA followed by Tukey-Kramer test, p˂0.0001). We thus elucidate a reduction in D-depth as a parameter characteristic of abnormal collagen fibrils in AngII-infused mice and human AAA.

Figure 7: Verification of abnormal collagen fibrils in AAA using atomic force microscopy (AFM).

AFM amplitude images show normal adventitial collagen fibrils with clearly resolvable D-periodic banding in AngII (B) and saline-infused mice (A), as well as in human AAA (E) and control tissue (D). Abnormal fibrils with weakened contrast in D-periodicity were also readily found in the aortic tissue from AngII-treated mice (C) and in human AAA (F). In certain regions fibrils with barely resolvable D-periodicity (white arrows) were found interspersed between normal fibrils (black arrows). Scale bars are 200 nm. AFM images of abnormal collagen fibrils from additional samples are shown in Figure S5.

Figure 8: Characterization of depth of D-periods in collagen fibrils from murine aorta.

AFM height images of normal fibrils and their corresponding section profiles for adventitial collagen in AngII-(C, D) and saline-infused mice (A, B) reveal a similar depth of D-period. However, abnormal collagen fibrils found in AngII-treated mice exhibited decreased D-depths (E, F). Scale bars are 200 nm. Histogram shows the distribution of D-period depth from a representative mouse sample as indicated (G). Average depth of D-periods from all samples shows no significant differences between normal fibrils from each treatment group. However significantly reduced D-depth were observed for abnormal fibrils in AngII-infused mice (*p˂0.0001) (H).

Figure 9: Characterization of depth of D-periods in collagen fibrils in human AAA.

AFM height images of collagen fibrils and their corresponding section profiles are shown for normal fibrils from human control (A, B) and clinical AAA normal (C, D) and for abnormal fibrils in clinical AAA (E, F). Scale bars are 200 nm. Histogram shows the distribution of D-period depth from a representative human sample as indicated (G). Average depth of D-periods from all samples shows no significant differences between normal fibrils from control or AAA groups. However, significantly reduced D-depth were observed for abnormal fibrils in AAA (*p˂0.0001) (H).

4. DISCUSSION

In this study, we employ two independent ultrastructural microscopy techniques, namely TEM and AFM to elucidate the presence of abnormal collagen fibrils with compromised D-periodic banding in the remodeled regions of AAA in two different species (murine and human). We used AngII-infused ApoE^−/− mice as it is a widely accepted mouse model of AAA[39]. Analysis of murine samples further enabled us to delineate that these abnormal fibrils were distinct from adventitial collagen fibrils present in the abdominal aorta of saline-infused mice and in non-dilated thoracic aorta of AngII-infused mice and from medial collagen in the abdominal aorta of saline-infused mice and were only present in the ECM remodeling accompanying AAA. Although the diameter of the abnormal fibrils resembled normal adventitial fibrils, they were characterized by a loss of D-periodicity (resulting in decrease in the depth of D-periods) and an increase in the local fibril curvature (Table 1). While these features were present in both mice and human AAA, they were more pronounced in the human AAA. This could partly be explained by the fact that severity of AAA may not be as advanced in AngII-infused ApoE knockout mice model[40] as compared to clinical samples. Our results showing structural changes in collagen fibrils are consistent with a recent study which reported how retardation of collagen fibrils (measured using polarization microscopy) is reduced in AAA[23]. Retardation values are indicative of changes in the molecular alignment and composition of collagen fibrils.

Table 1:

Summary of measurements of various parameters in collagen fibrils in murine and human AAA.

		Depth of D-Period (nm)	Fibril Diameter (nm)	Fibril Curvature (1/μm)
Mouse	Saline (Normal)	3.3 ± 0.6 (6; 224)	53 ± 3 (6; 2269)	1.0 ± 0.3 (6; 60)
	Angll-TA (Normal)	3.2 ± 0.2 (3; 57)	57 ± 4 (3; 540)	1.3 ± 0.2 (3; 30)
	Angll (Normal)	3.3 ± 0.3 (8; 111)	53 ± 6 (6; 1538)	0.9 ± 0.2 (6; 60)
	Angll (Abnormal)	0.9 ± 0.1 (8; 267)	55 ±3(6; 3459)	3.1 ± 0.5 (6; 60)
Human	Control (Normal)	4.0 ± 0.3 (6; 354)	47 ± 11 (6; 930)	1.4 ± 0.1 (6; 60)
	AAA (Normal)	3.8 ± 0.3 (6; 163)	45 ± 9 (6; 978)	2.1 ± 0.5 (6; 60)
	AAA (Abnormal)	1.0 ± 0.1 (6; 213)	38 ± 9 (6; 1011)	4.3 ± 0.7 (6; 60)

Means ± SD are listed with number of mice (n_m) and number of collagen fibrils (n_f), or number of D-periods (n_p) in parentheses (n_m; n_f or n_p).

Shaded cells: means are statistically significant (p < 0.0001).

The loss of D-periodicity in abnormal fibrils in AAA as shown in our study can be attributed to multiple factors. In particular during collagen synthesis, a dysregulated collagen fibrillogenesis could lead to alterations in the molecular packing of collagen fibrils resulting in structural abnormalities. ECM remodeling in AAA is characterized by synthesis of new collagen as an important compensatory mechanism. We and others have shown that the two major cell types dictating collagen synthesis in aneurysmal tissue are synthetic SMCs[18][34][15] (trans-differentiated from the contractile SMC phenotype present in the media) and myofibroblasts[41][42] (trans-differentiated from adventitial fibroblasts). Although the gene expression and regulation of collagen synthesis by these cells[1][43] is extensively characterized in cardiovascular pathologies, very little is understood about the regulation of collagen fibrillogenesis and fibril structure by these trans-differentiated cells. It is interesting to note that collagen fibrillogenesis is modulated by a multitude of factors including matricellular proteins (e.g. fibromodulin[44] and decorin[45]), collagen receptors[38][46,47], other collagen types[48] and crosslinking enzymes (e.g. lysyl oxidase[49]), several of which have been reported upregulated in AAA. It is thus reasonable to surmise that both the cell type and the biochemical composition of the local ECM environment may play a role in synthesis of abnormal collagen fibrils.

Degradation of pre-existing normal collagen fibrils also result in morphological changes. Several collagenases have been reported to be upregulated in AAA tissues[50]. However, recent structural characterization of collagen fibrils from rat tendon-fascicles has revealed that after the action of the matrix metalloproteases (MMP1 and MMP13), the collagen fibril is broken up into smaller structural units which still preserve the native D-periodicity [51]. Thus even though MMPs are an unlikely contributor for abnormal collagen fibrils, there is some evidence that other collagenases such as Cathepsin K may result in a loss of D-periodicity[52] and elastases such as Cathepsin S contribute to collagen degradation[53]. It should be noted that the synthesis and degradation of collagen fibrils may also be affected by inflammatory factors present in AAA[9] as chronic inflammation in temporomandibular joints has recently been reported to result in undulating collagen fibrils with diminished depth of D-periods[54].

Mechanical insult can thus be another putative cause of abnormal collagen fibrils in AAA. Ex-vivo studies on healthy aorta have elucidated how removal of elastin results in similar structural changes in the collagen fibers as in mechanical deformation[55]. A number of experimental and computational studies in AAA have elucidated how micro-structural damage co-localizes with excessive mechanical strain[56,57]. In particular the disruption of elastic fibers in AAA leaves collagen fibers as the main load-bearing agent, which would experience increased compression in the circumferential direction and increased stretch in the axial direction with the growing aneurysm [9]. In this regard, over-stretching of cerebral arteries ex-vivo was shown to result in collagen damage which could be detected using a collagen hybridizing peptide[58]. At the ultrastructural level, a fuzzy surface texture, obscuring or completely hiding the D-periodic banding in collagen fibrils has been reported for mechanically overloaded rat-tendon collagen fibrils, a phenomenon termed discrete plasticity[59]. X-ray diffraction studies on corneal collagen with applied tensile load in-situ has also revealed small changes in the length of D-periods and molecular orientation within the collagen fibril[60].

Mechanical loading and mechanical properties can also be attributed to the increased curvature or undulations present in abnormal collagen fibrils. Studies from other tissues have postulated that undulating fibrils are better suited to respond to multidirectional loadings[61][62]. One may thus speculate that the abnormal collagen in AAA represents newly synthesized collagen in response to redistribution of mechanical load in ECM remodeling. Alternatively a perturbation in collagen fibrillogenesis can also result in collagen fibrils with increased curvature as shown in our earlier study[63].As another paradigm, mechanical loading may perturb the structure of preexisting fibrils. In this regard, ex-vivo cyclic loading of tendon tissue has been shown to result in fatigue damage (manifested as kinks) in a subset of collagen fibrils[64]. Thus multiple factors including mechanical load, dysregulation in synthesis and degradation of collagen and presence of a pro-inflammatory environment in ECM remodeling in AAA may be causal to abnormal fibrils.

Abnormal collagen fibrils could potentially impact the functional properties of the AAA. In this regard, it is interesting to note that AFM nano-indentation measurements of AAA tissue has revealed regions with drastically different stiffness values[4]. Our studies suggest that this heterogeneity in stiffness may correspond to regions of normal versus abnormal fibrils. Secondly, the fibril structure can also affect their susceptibility to degradation by gelatinases (e.g. MMP9) as shown for mechanically damaged fibrils in other tissues[65]. Such a scenario would be particularly relevant for AAA, as increased MMP9 activity has been observed in local spots in the aneurysm tissue using in-situ zymography[66] and an increased MMP9 level correlated to AAA rupture[67]. Besides local mechanical heterogeneity and an enhanced ability for degradation, abnormal collagen fibrils may also impact cell-matrix interactions by modulating the exposure or accessibility of specific epitopes on the fibril surface[68]. For instance collagenase degraded fibrils have been elucidated to alter cell motility, signaling and gene expression[51] and it is possible that abnormal fibrils may likewise differentially regulate cell-behavior as compared to normal fibrils.

Taken together, our investigations have identified and characterized abnormal collagen fibrils with two quantitative metrics (changes in fibril curvature and depth of D-periods) as a distinct feature of pathological ECM remodeling accompanying AAA. Our results can serve as a foundation for further studies to establish abnormal collagen fibrils as a hallmark of ECM remodeling in AAA and potentially in other cardiovascular pathologies. Further experiments designed to characterize the causative factors, biochemical signatures, temporal and spatial distribution and functional properties of structurally altered abnormal collagen fibrils may provide further insights into their role in vascular diseases. In addition, abnormal collagen fibrils could potentially be targeted via specific contrast agents to characterize dysfunctional matrix turnover in AAA, in a manner similar to that being pursued for accumulation of monomeric tropoelastin[69].

Statement of Significance.

Several vascular diseases including abdominal aortic aneurysm (AAA) are characterized by extensive remodeling in the vessel wall. However, although structural alterations in elastin fibers are well characterized in vascular diseases, very little is known about the collagen fibril structure in these diseases. We report here a comprehensive ultrastructural evaluation of the collagen fibrils in a mouse model of AAA, using high-resolution microscopy techniques like transmission electron microscopy (TEM) and atomic force microscopy (AFM). We elucidate how abnormal collagen fibrils with compromised D-periodicity and increased fibril curvature are present in the vascular tissue in both clinical AAA as well as in murine models. We discuss how these abnormal collagen fibrils are likely a consequence of mechanical overload accompanying AAA and could impact the functional properties of the underlying tissue.

Abbreviations:

AAA

Abdominal aortic aneurysm

AFM

atomic force microscopy

AngII

angiotensin II

EDS

Ehlers Danlos Syndrome

ECM

extracellular matrix

MMP

matrix metalloprotease

sCAD

spontaneous cervical artery dissection

SMC

smooth muscle cell

TEM

transmission electron microscopy