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Ann Thorac Surg 2007;83:1641-1650
© 2007 The Society of Thoracic Surgeons

Original Articles: Cardiovascular

Optimized Preservation of Extracellular Matrix in Cardiac Tissues: Implications for Long-Term Graft Durability

Katja Schenke-Layland, PhDa, Jiansong Xie, MD, PhDb, Sepideh Heydarkhan-Hagvall, PhDc, Sarah F. Hamm-Alvarez, PhDb, Ulrich A. Stock, MDd, Kelvin G.M. Brockbank, PhDe,f, W. Robb MacLellan, MDa,*

a Cardiovascular Research Laboratory, David Geffen School of Medicine at UCLA, Los Angeles, California
b Department of Pharmaceutical Sciences, School of Pharmacy, USC, Los Angeles, California
c Regenerative Bioengineering and Repair Laboratory, UCLA, Los Angeles, California
d Department of Medical Physics and Biophysics, Humboldt University, University Hospital Charité, Berlin, Germany
e Cell & Tissue Systems, Inc., Charleston, South Carolina
f The Georgia Tech/Emory Center for The Engineering of Living Tissues, Georgia Institute of Technology, Atlanta, Georgia

Accepted for publication December 4, 2006.

* Address correspondence to Dr MacLellan, University of California Los Angeles, Cardiovascular Research Laboratory, 675 Charles E. Young Drive South, MRL 3-645, Los Angeles, CA 90095-1760 (Email: rmaclellan{at}mednet.ucla.edu).


Dr Brockbank discloses that he has a financial relationship with Cell and Tissue Systems, Inc.

 

   Abstract
Top
 Abstract
Introduction
 Material and Methods
 Results
 Comment
 Acknowledgments
 References
 
Background: Cryopreservation of human tissues, particularly heart valves, is widespread in clinical practice although the effects of this process on underlying tissue structures and its potential impact on valve durability have been poorly studied. Multiphoton imaging and second-harmonic generation (SHG) microscopy permit high-resolution, noninvasive analysis of living tissues at a subcellular level. In the present study we used these novel imaging modalities to compare the effects of vitreous and frozen cryopreservation on the extracellular matrix (ECM) of cardiac tissues.

Methods: Conventional histology, electron microscopy, and multiphoton imaging to obtain autofluorescence and SHG images were performed on cardiac tissues to characterize the ECM in fresh, vitrified, and frozen cryopreserved tissues.

Results: Autofluorescence and particularly SHG images revealed that conventional frozen cryopreservation of cardiac valves, when compared with fresh or vitrified tissues, leads to the loss of normal ECM structures in valve leaflets. Similar results were found in all other cardiac tissues suggesting that structural deterioration of the ECM is a common consequence of frozen cryopreservation.

Conclusions: Our results demonstrate that conventional cryopreservation, when compared with fresh or vitrified tissues, causes more destruction of normal ECM structure, which might contribute to eventual graft dysfunction. Whether vitrification preservation will translate into greater durability or less valve failure will need to be determined.

The necessity for human tissues and organs for reconstructive medicine is driving a search for novel and more advanced tissue storage and biopreservation technologies. Despite current research efforts, little is known regarding the pathophysiology of graft failure, particularly regarding the role that the preservation technique itself has on cellular viability, immune responses, and especially extracellular matrix (ECM) structure. Identifying and quantifying the multifactorial mechanisms that lead to failure of cryopreserved cardiac tissues is therefore an important clinically relevant issue that will need to be addressed for the field to progress.

Currently, there are several approaches to the preservation and storage of living human biomaterials [1–3], but cryopreservation by controlled freezing using cryoprotectant solutions is the primary preservation method for cardiovascular tissue banking. However, this technique has a number of limitations and does not adequately preserve complex multicellular tissues, such as heart valves or blood vessels. Frozen cryopreserved cardiac tissues display both reversible and irreversible cell injury, including destruction of the ECM with poor cell viability and functionality [4–6]. The damage induced by cryopreservation has in large part been ascribed to the phenomenon of interstitial ice formation [7, 8]. Indeed, cryopreservation-induced ECM injury may predispose valvular grafts to calcification in vivo, resulting in the early structural deterioration and dysfunction of cryopreserved allogeneic homografts [9]. Vitrification is an alternative approach to conventional long-term subzero preservation, which may overcome some of these limitations. Vitreous cryopreservation uses high concentrations of a cryoprotectant solution to promote amorphous solidification rather than crystallization to restrict the amount of ice crystal formation [3, 10]. However, whether this theoretical advantage translates into less damage of preserved tissues is unknown.

In this study we assessed the impact of conventional cryopreservation versus vitrification on state and quality of ECM structures (such as collagenous bundles and elastin-containing fibers) in a variety of cardiac tissues including aortic and pulmonary cardiac muscle, blood vessels, and heart valve leaflets, using conventional histologic and electron microscopy analysis, as well as in situ multiphoton and second harmonic generation (SHG) imaging. Multiphoton-induced autofluorescence excitation based on the simultaneous absorption of two or more near-infrared photons enables the nondestructive deep-tissue imaging of endogenous fluorophores, which allows an artifact-free, nondestructive tissue state evaluation of unprocessed tissues and organs [11, 12].

Our data demonstrate that conventional cryopreservation leads to a loss of normal ECM structure, suggesting that vitrification may be a promising alternative approach for cardiac tissue preservation that preserves ECM architecture.


   Material and Methods
Top
 Abstract
 Introduction
 Material and Methods
Results
 Comment
 Acknowledgments
 References
 
Sample Preparation
Porcine hearts were obtained from a local slaughterhouse. Aortic and pulmonary valves were excised under sterile conditions and incubated in an antibiotic solution consisting of Dulbecco’s modified Eagle medium (DMEM, MediaTech, Herndon, VA), containing 4.5 g/L glucose and 1% penicillin-streptomycin (Sigma, St. Louis, MO) at 4°C prior to use. All valves were further processed according to established frozen cryopreservation and vitrification protocols. For conventional cryopreservation by freezing, the valves were processed according to the steps disclosed in U.S. Patent 4,890,457 [13], followed by thawing in two stages. In step 1 the tissues were slowly warmed to –100°C by placement in a cooler with dry ice without direct contact to the latter to avoid freeze damage. In step 2 they were rapidly warmed in a 37°C water bath until all ice had visibly disappeared. This warming protocol follows current clinical practice.

Vitreous Cryopreservation
Ice-free cryopreservation by vitrification was achieved by gradually infiltrating the heart valve tissues in six 15-minute-steps at 4°C, with increasing concentrations of a precooled vitrification solution to achieve a final concentration of 8.4 mol/L (VS55; made up of three components: 3.10 mol/L DMSO, 3.10 mol/L formamide and 2.21 mol/L 1,2-propanediol [all Sigma] in Euro-Collins solution) [14]. After the final step the heart valves were placed individually in sterile polyester bags containing 80 mL of the vitrification solution. A dummy sample was established with a built-in, nonintrusive thermocouple in the center of the heart valve-bag attached to a digital thermometer to monitor experimental temperatures during cryopreservation. Each bag was evacuated of air, heat-sealed, and cooled at 5.4°C/minute to –100°C in a precooled bath of 2-methylbutane (isopentane). The bags were then transferred to the top (vapor phase) of a liquid nitrogen storage freezer and slowly cooled at greater than 0.5°C/minute to –135°C. The valves were stored at –150°C for a minimum of 24 hours prior to thawing for study. The vitrified valves were rewarmed using a two-step process including slow warming to –100°C using convection at 8.6°C/minute, followed by faster warming to –35°C at 17°C/minute in a water bath at 37°C. The concentration of the vitrification solution was then decreased (cryoprotectants eluted) from 8.4 mol/L vitrification solution to DMEM containing 10% fetal bovine serum in seven steps on ice. All samples were stored at 4°C in DMEM until processed.

Routine Histology and Transmission Electron Microscopy
For routine histological analysis, heart valves were processed as previously described [11]. A modified Movat-pentachrome stain [15] was used to demonstrate ECM components. For improved histologic imaging of elastic fibers, tissue slides were stained with a modified Hart’s resorcin-fuchsin stain [16]. For transmission electron microscopy (TEM) heart valves were processed and analyzed as previously described [17].

Multiphoton Imaging and SHG Microscopy
Multiphoton imaging and SHG microscopy were performed using a Zeiss LSM 510 META NLO femtosecond laser scanning system (Carl Zeiss), coupled to a software-tunable Coherent Chameleon titanium:sapphire laser (720 nm to 930 nm, 90 MHz; Coherent Laser Group, Santa Clara, CA), and equipped with a high-resolution AxioCam HRc camera with 1300 x 1030 pixels (Carl Zeiss). Images were collected using an oil immersion Plan-Neofluar 40x/1.3 numerical aperture (NA) differential interference contrast, or an oil immersion Plan Apochromat 63x/1.4 NA objective lens (Carl Zeiss). All observations were made using unprocessed, untreated intact tissues. The ECM structure-dependent autofluorescence and SHG were induced using wavelengths of 760 nm (elastin) and 840 nm (collagen) as described previously in more detail [11, 17, 18]. Noninvasive serial optical horizontal sections of four different areas of each of the specimens were taken in z-steps of 1 µm, 2.5 µm, or 5 µm to depths of 50 to 100 µm.

SHG Signal Profiling by Spectral Fingerprinting and Statistical Data Analysis
To quantify the intrinsic fluorescence signals of collagenous structures within fresh, vitrified, and frozen cryopreserved cardiac tissues, lambda stacks were ascertained at emission wavelengths of 392 nm to 499 nm (in 10 nm increments), using the two-photon Chameleon laser tuned to an excitation wavelength of 840 nm. Emission was collected using the Zeiss META detector (spectral separator) of the LSM 510 Meta NLO system. The scans were acquired with a scan time of 2.56 µs/pixel, 8 times summarization to enhance the signal, in plane multitrack 8-bit lambda mode, and analyzed by the Zeiss AIM version 3.2 software (Carl Zeiss). Due to a reproducible laser excitation power and similar exposure times for all samples, mean intensities of the intrinsic SHG signals were calculated and used for quantification. Briefly, in each case four different areas of each tissue sample were scanned and intensities of the intrinsic SHG signals of the regions of interest (ROI; equals the area of the highest intensity) were detected. The SHG signal intensities were reflected by the gray values of all the pixels within a ROI. Mean intensities were calculated from the four screened ROI areas. Accordingly, all results are presented as mean values ± standard deviations (SD). Significant differences between fresh, vitrified, and frozen cryopreserved tissues were assessed by analysis of variance with the Tukey multiple comparison test. The p values of 0.05 or less were defined as statistically significant.


   Results
Top
 Abstract
 Introduction
 Material and Methods
 Results
Comment
 Acknowledgments
 References
 
Morphological and Microscopic Analysis of Cryopreserved Cardiac Tissue
Macroscopic visualization did not reveal differences between fresh unprocessed, thawed vitrified, and thawed frozen cryopreserved valves. To determine if changes were present at the microscopic level, routine histology was performed (Fig 1). Compared with fresh specimens (Figs 1A, 1D, 1G), the structure of cardiac muscle, trunk, and leaflets of rewarmed vitrified tissues (Figs 1B, 1E, 1H) demonstrated comparable morphologic features with minimal ECM changes. In contrast, the ECM of thawed frozen cryopreserved samples (Figs 1C, 1F, 1I) appeared to be more fragmented and less organized. Compared with the histoarchitecture of fresh cardiac muscle, frozen cryopreserved specimens showed a looser appearance of the ECM, including large disrupted spaces in between the interwoven network of cardiac muscle cells (Figs 1A vs 1C). Moreover, microscopy of Hart’s stained frozen cryopreserved trunk sections revealed a greater degree of elastic fiber fragmentation, characterized by disrupted fibers and increased interfibrillar spaces (Figs 1D vs 1F). The ECM of vitrified and frozen cryopreserved heart valve leaflets showed a preserved layered leaflet structure; however, within the inner spongiosa layer of the leaflets structural imperfections were visible in frozen cryopreserved heart valve leaflets and to a lesser extent in vitrified valve leaflets (Figs 1G through 1I).


Figure 1
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  		Fig 1. Representative light-micrographs of fresh (A, D, G), thawed vitrified (B, E, H), and thawed frozen cryopreserved (C, F, I) aortic cardiac tissues. (A–C) Movat-pentachrome stained cross-sections of cardiac muscle; (D–F) Hart’s stained slides of aortic vascular tissue; (G–I): of the inflow side (v, ventricularis), the inner layer (s, spongiosa), and the valve outflow side (f, fibrosa) of aortic heart valve leaflets. (A through F scale bar equals 200 µm; G through I scale bar equals 50 µm.)

 
Ultrastructural Analysis of Valvular Tissues
To initially analyze the ultrastructure of fresh, vitrified, and frozen cryopreserved heart valves we performed conventional TEM, showing relatively few changes in the cellular and extracellular components of vitrified and frozen cryopreserved tissues compared with fresh valve leaflets (Fig 2). However, a closer examination of the inflow sides of vitrified and frozen cryopreserved heart valve leaflets revealed an overall loss of the elastin-associated microfibrillar network (Figs 2B, 2C). These changes were also seen in other elastin-rich tissues, such as aortic and pulmonary trunk (data not shown).


Figure 2
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  		Fig 2. Electron micrographs of representative parts of the ventricularis layer of fresh (A), vitrified (B), and frozen cryopreserved (C) pulmonary heart valve leaflets, demonstrating elastic fibers (EF), including microfibrills (MF), and collagenous fibers (CF). Notice that within vitrified as well as frozen cryopreserved tissues almost no microfibrillar components could be detected.

 
To confirm that these changes were a direct result of the cryopreservation technique, and did not occur secondary to the fixation required for TEM scanning, we screened nonprocessed (nonfixed, nonsliced, nonstained) fresh, vitrified, and frozen cryopreserved heart valve tissues using multiphoton-excited autofluorescence and SHG microscopy (Figs 3; 4; 5). Go Go Using this imaging modality, ECM components such as collagen and elastic fibers can be nondestructively detected at two different laser excitation wavelengths. Exposure to laser pulses at 760 nm revealed single cells with fluorescent mitochondria based on two-photon excited reduced coenzyme NAD(P)H (predominantly visible within leaflet tissues), and elastic fibers (green structures; Figs 3; 4; 5A, 5C, 5E, 5G, 5I, 5K). Imaging of the same intratissue regions at a higher wavelength (840 nm) showed wavelike or dense bundled collagenous fibers (red; Figs 3; 4; 5A, 5C; 5E, 5G; 5I, 5K). Due to femtosecond laser-induced SHG processes, these structures emitted blue light at the peak emission wavelength of 413 nm, which served as a quantitative measure of structural preservation or damage of vitrified and frozen cryopreserved tissues (Figs. 3; 4; 5B, 5D; 5F, 5H; 5J, 5L; Table 1).


Figure 3
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  		Fig 3. Multiphoton-induced autofluorescence imaging and second-harmonic generation (SHG) signal profiling of the collagen-rich outflow (A,B; E,F; I,J) and the predominantly elastin-containing inflow side (C,D; G,H; K,L) of representative areas of fresh (A–D), vitrified (E–H) and frozen cryopreserved (I–L) aortic leaflets. The graphs represent peak SHG intensities of collagen-containing structures in the corresponding lambda stack overlay images, denoted by the red cross (B,D; F,H; J,L). The SHG imaging demonstrates the substantial ultrastructural deterioration and disintegration of most collagenous structures in cryopreserved tissues (J, L). Sufficient SHG signals were only inducible in fresh and vitrified tissue structures (B,D; F,H). Compared with fresh and vitrified tissues (A,C; E,G), just a few elastic fibers are detectable within cryopreserved leaflets (I, K). Living cells were only found within fresh and some of the vitrified leaflet tissues (white arrows, E).

 

Figure 4
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  		Fig 4. Multiphoton imaging (A,C; E,G; I,K) and second-harmonic generation profiling (B,D; F,H; J,L) of aortic (A,B; E,F; I,J) and pulmonary (C,D; G,H; K,L) cardiac muscle specimens. Collagenous structures (red: 840 nm), elastic fibers and cells (green: 760 nm) are clearly visible within fresh (A–D) and vitrified (E–H) tissues. Only weak autofluorescence signals are detectable in frozen cryopreserved cardiac muscle (I–L).

 

Figure 5
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  		Fig 5. Two-photon images of aortic (A, E, I) and pulmonary (C, G, K) trunk regions induced with wavelengths of 760 nm (green: elastic fibers, cells) and 840 nm (red: collagen). In comparison with the fresh controls (A, C), vascular vitrified structures (E, G) are well preserved, whereas the ECM of frozen cryopreserved tissues (I, K) appears to be damaged and not well organized. The SHG signal detection of the collagenous fibers of aortic (B, F, J) and pulmonary (D, H, L) trunk regions by spectral fingerprinting reveals a dramatic loss of signal intensities, especially within the pulmonary frozen cryopreserved specimens (L) when compared with fresh (D) tissues.

 

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  		Table 1 Summary of SHG Signal Intensities of Collagenous Structures Detected by Spectral Fingerprinting. All Sample Data are Shown as Mean ± SD of the Maximal Intensity of the Emission Value Detected at the Peak Emission Wavelength of 413 nm
 
Multiphoton imaging demonstrated branched elastic fibers and wavy bundles of collagen in all fresh and vitrified heart valve leaflets (Figs 3A, 3C; 3E, 3G). Moreover, single living cells were visible within some of the vitrified samples (Fig 3E). Although SHG signals of the vitrified tissues were weaker when compared with the fresh specimens (Fig. 3B,3D vs 3F,3H; Table 1), almost no autofluorescence and SHG signals were inducible within frozen cryopreserved specimens (Figs 3I to 3L; Table 1). When collagen fibers lose their native structure and density they generate less detectable SHG signals. Thus, the diminished SHG signal intensities are indicative of structural changes of the collagenous fibers in the analyzed valve tissues. The weaker the SHG signals, the more changes of the ECM structures occurred. To quantify these changes we detected SHG signal intensities of collagenous structures (Table 1). In comparison with the fresh controls, vitrified and frozen cryopreserved aortic heart valve leaflet samples showed a 44% and 77% decrease in SHG intensities, respectively. Similar reductions were seen in vitrified versus frozen cryopreserved pulmonary leaflets. This decrease in intrinsic fluorescence signals of collagenous structures in frozen cryopreserved specimen was significantly greater than in vitrified tissues (p < 0.001; in both aortic and pulmonary tissues).

Multiphoton and SHG Imaging of Cardiac Muscle and Vascular Tissues
To determine if the damaging effects of cryopreservation on ECM occurred in all tissues we examined cardiac muscle and vascular tissues from the same valve specimens (Figs 4; 5; Table 1). Similar to leaflet tissues, cellular structures, elastic fibers, and wavy bundles of collagen were detectable in all fresh and thawed vitrified aortic cardiac muscle specimens (Figs 4A, 4E). Likewise, there was a dramatic loss of autofluorescence and SHG within all thawed frozen cryopreserved samples (Figs 4I to 4L; Table 1). Using the same laser power as for the detection of the aortic tissues, all pulmonary cardiac muscle specimens showed reduced SHG intensities (Figs 4D, 4H, 4L; Table 1). A significant loss of autofluorescence and SHG was detectable within pulmonary vitrified and frozen cryopreserved samples (Figs 4G, 4H; 4K, 4L; Table 1).

Imaging of fresh and thawed vitrified aortic or pulmonary trunk regions showed a similar pattern of elastin-containing structures (Figs 5A, 5C; 5E, 5G). However, SHG signals of collagenous fibers of the pulmonary vitrified samples were significantly weaker than those of the fresh controls (Figs 5B, 5D vs 5F, 5H; Table 1). Poor autofluorescence and almost no SHG signals were detectable within aortic or pulmonary frozen cryopreserved specimens (Figs 5I to 5L; Table 1).


   Comment
Top
 Abstract
 Introduction
 Material and Methods
 Results
 Comment
Acknowledgments
 References
 
To date, the effects of vitreous or conventional frozen cryopreservation on the quality and integrity of ECM components in preserved cardiovascular tissues are largely unknown. Currently, the method of choice for the long-term storage of tissue grafts, such as heart valves or blood vessels, has been frozen-preservation at ultralow temperatures, although it is known that damage of extracellular tissue structures by the crystallization of ice during the freeze-thaw cycle is a major obstacle to graft preservation, which consequently affects their long-term performance after implantation [19, 20]. The data from this study suggest that ice-free vitreous cryopreservation may overcome, at least some of the limitations of current preservation techniques. Vitrification has been shown to provide effective preservation for a number of tissues including oocytes, early embryos, cartilage, and skin [21–24]. Furthermore, vitrified arterial blood vessels and tissue-engineered vascular grafts have shown to maintain viscoelastic properties similar to fresh vascular tissues, and demonstrated superior biomechanical performance when compared with frozen cryopreserved specimens [25–27].

In previous studies, ECM properties of vitrified or frozen cryopreserved tissues were assessed either by functional biomechanical tests or by relatively insensitive techniques including histologic staining, electron microscopy, or biochemical analysis. While these methods are useful tools for general matrix characterization, they are inadequate to predicate the state of ECM structures within a tissue. In this study, conventional histologic staining using Movat-pentachrome and a modified Hart’s resorcin-fuchsin stain revealed almost comparable cell and ECM formations within fresh, vitrified, and frozen cryopreserved aortic and pulmonary cardiac muscle, trunk, and valve leaflet tissues. The histoarchitecture of vitrified and frozen cryopreserved specimens appeared only mildly disturbed, with a looser appearance of the ECM and some structural imperfections within the layered structure of the leaflets, as well as partial elastic fiber fragmentation, particularly in frozen cryopreserved trunk tissues. Although electron microscopy showed minimal loss of the elastin-associated microfibrillar network in both vitrified and frozen cryopreserved tissues, this conventional technique dramatically underestimated the impact of cryopreservation on essential matrix structures. Significant changes of the ECM were only revealed using multiphoton-excited autofluorescence and SHG microscopy. Thus, routine methodologies are not sufficient if a detailed tissue state characterization is required.

Multiphoton-excited autofluorescence and SHG microscopy are powerful imaging tools that provide the unique ability to detect deep-tissue cells and ECM components such as elastic and collagenous fibers in situ without the need for any fixation or invasive processing [12, 28]. In particular, the fluorescent coenzyme NAD(P)H, flavoproteins, keratin, melanin, and elastin are easily detected by two-photon excited autofluorescence using different excitation wavelengths [12]. Myosin, tubulin, and the ECM protein collagen can also be imaged by the second-order nonlinear optical effect—the SHG process [29]. In our study, multiphoton imaging and SHG microscopy revealed only weak autofluorescent structures within the cryopreserved samples when exposed to laser pulses at 760 nm and especially at 840 nm. The SHG signals were almost not detectable. In contrast, imaging of vitrified cardiac specimens demonstrated better preservation of ECM structures, comparable with fresh tissues. Those imaging results were strengthened by the assessment of intrinsic SHG signals using spectral fingerprinting, which served as a quantitative measure of structural preservation or damage of intratissue collagenous fibers. Thus, although both preservation methods caused changes to the native properties of ECM structures, when compared with vitrification, cryopreservation by controlled freezing had a significantly higher impact on the integrity of ECM components.

The direct impact of the loss of normal ECM pattern on long-term graft durability and survival is not well understood. However, extracellular structures including the elastin-associated microfibrillar network are crucial for elastic fiber assembly, are directly associated with nonfibrillar (glycosaminoglycans) as well as fibrillar ECM components and thus are predicted to play an important role in matrix structure maintenance [30]. Several reports have suggested that there is a direct correlation between general ECM degeneration and destructive calcification of heart valve grafts [9, 31]. Moreover, evidence exists that disorganization of collagenous fibers, and in particular the destruction of the elastic fiber network, leads to medial vascular calcification. Similar processes might be involved in pathological heart valve calcification [32, 33]. However, the exact mechanisms that are responsible for calcific heart valve degeneration, leading to graft dysfunction, are still unclear. Further in vitro and in vivo studies focusing on the role of ECM proteins such as elastin or tenascin-C [34], matrix metalloproteinases and inflammatory processes are necessary.

In conclusion, cryopreserved homograft or allograft valves are well-established valvular substitutes. In the majority of cases they show good hemodynamic properties, are resistant to infections, and exhibit a low incidence of thromboembolic complications; however, their long-term durability has been questioned, especially in children, young adults, and patients with terminal renal failure [35]. The present results demonstrate that both conventional freezing and vitrification preservation have limitations and lead to changes in the ECM tissue architecture. However, in this study vitrification led to less ECM damage within the cardiac tissues analyzed than conventional cryopreservation. Whether this will translate into greater long-term durability or less valve failure will need to be determined, but given the current data suggesting a link between ECM damage and graft calcification, vitrification may represent a clinical advancement in tissue transplant and reconstructive therapies.


   Acknowledgments
Top
 Abstract
 Introduction
 Material and Methods
 Results
 Comment
 Acknowledgments
References
 
The authors would like to thank Sabine Hitschke for her excellent technical assistance. We are grateful to Prof Dr Karsten König and Dr Ekaterini Angelis for their helpful discussions and thoughts. This work was supported by gifts from the Laubisch Fund as well as the Deutsche Forschungsgemeinschaft - DFG (Sche701/2-1, 3-1) (K.S-L.), NIH EY-11386 (S.H-A.), NIH R01 HL 70748, P01 HL080111, and AHA 0340087N grants (W.R.M.).


   References
Top
 Abstract
 Introduction
 Material and Methods
 Results
 Comment
 Acknowledgments
 References
 

  1. Hicks M, Hing A, Gao L, Ryan J, Macdonald PS. Organ preservation Methods Mol Biol 2006;333:331-374.[Medline]
  2. Fahy GM, Wowk B, Wu J. Cryopreservation of complex systems: the missing link in the regenerative medicine supply chain Rejuvenation Res 2006;9:279-291.[Medline]
  3. Brockbank KG, Taylor MJ. Tissue preservationIn: Baust JG, Baust JM, editors. Advances in Biopreservation. Boca Raton, FL: CRC Press/Taylor & Francis Group; 2006. pp. 157-192.
  4. Messier RH, Bass BL, Domkowski PW, Hopkins RA. Interstitial cellular and matrix restoration of cardiac valves after cryopreservation J Thorac Cardiovasc Surg 1999;118:36-49.[Abstract/Free Full Text]
  5. Villalba R, Pena J, Luque E, Gomez Villagran JL. Characterization of ultrastructural damage of valves cryopreserved under standard conditions Cryobiology 2001;43:81-84.[Medline]
  6. Kitagawa T, Masuda Y, Tominaga T, Kano M. Cellular biology of cryopreserved allograft valves J Med Invest 2001;48:123-132.[Medline]
  7. Brockbank KG, Lightfoot FG, Song YC, Taylor MJ. Interstitial ice formation in cryopreserved homografts: a possible cause of tissue deterioration and calcification in vivo J Heart Valve Dis 2000;9:200-206.[Medline]
  8. Brockbank KG, Song YC. Morphological analyses of ice-free and frozen cryopreserved heart valve explants J Heart Valve Dis 2004;13:297-301.[Medline]
  9. Brockbank KG, Song YC. Mechanisms of bioprosthetic heart valve calcification Transplantation 2003;75:1133-1135.[Medline]
  10. Fahy GM, Wowk B, Wu J, et al. Cryopreservation of organs by vitrification: perspectives and recent advances Cryobiology 2004;48:157-178.[Medline]
  11. Schenke-Layland K, Madershahian N, Riemann I, et al. Impact of cryopreservation on extracellular matrix structures of heart valve leaflets Ann Thorac Surg 2006;81:918-926.[Abstract/Free Full Text]
  12. Schenke-Layland K, Riemann I, Damour O, Stock UA, König K. Two-photon microscopes and in vivo multiphoton tomographs – Powerful diagnostic tools for tissue engineering and drug delivery Adv Drug Deliv Rev 2006;58:878-896.[Medline]
  13. McNally RT, Heacox A, Brockbank KGM, Bank HL. Method for cryopreserving heart valves. US Patent #4,890,457 (1990).
  14. Khirabadi BS, Song YC, Brockbank KGM. Method of cryopreservation of tissues by vitrification. US Patent #6,740,484 (2004).
  15. Russell Jr HK. A modification of Movat’s pentachrome stain Arch Pathol 1972;94:187-191.[Medline]
  16. Starcher B, Aycock RL, Hill CH. Multiple roles for elastic fibers in the skin J Histochem Cytochem 2005;53:431-443.[Abstract/Free Full Text]
  17. Schenke-Layland K, Riemann I, Stock UA, König K. Imaging of cardiovascular structures using NIR femtosecond multiphoton laser scanning microscopy J Biomed Opt 2005;10:024017.[Medline]
  18. Konig K, Riemann I. High-resolution multiphoton tomography of human skin with subcellular spatial resolution and picosecond time resolution J Biomed Opt 2003;8:432-439.[Medline]
  19. Brockbank KG, Song YC, Khirabadi BS, Lightfoot FG, Boggs JM, Taylor MJ. Storage of tissues by vitrification Transplant Proc 2000;32:3-4.[Medline]
  20. Wusteman MC, Pegg DE, Robinson MP, Wang LH, Fitch P. Vitrification media: toxicity, permeability, and dielectric properties Cryobiology 2002;44:24-37.[Medline]
  21. Oktay K, Cil AP, Bang H. Efficiency of oocyte cryopreservation: a meta-analysis Fertil Steril 2006;86:70-80.[Medline]
  22. Boonkusol D, Gal AB, Bodo S, Gorhony B, Kitiyanant Y, Dinnyes A. Gene expression profiles and in vitro development following vitrification of pronuclear and 8-cell stage mouse embryos Mol Reprod Dev 2006;73:700-708.[Medline]
  23. Song YC, An YH, Kang QK, et al. Vitreous preservation of articular cartilage grafts J Invest Surg 2004;17:65-70.[Medline]
  24. Silvestre MA, Sanchez JP, Gomez EA. Vitrification of goat, sheep, and cattle skin samples from whole ear extirpated after death and maintained at different storage times and temperatures Cryobiology 2004;49:221-229.[Medline]
  25. Song YC, Khirabadi BS, Lightfoot FG, Brockbank KGM, Taylor MJ. Vitreous cryopreservation maintains the function of vascular grafts Nature Biotech 2000;18:296-299.[Medline]
  26. Dahl SL, Chen Z, Solan AK, Brockbank KG, Niklason LE, Song YC. Feasibility of vitrification as a storage method for tissue-engineered blood vessels Tissue Eng 2006;12:291-300.[Medline]
  27. Thakrar RR, Patel VP, Hamilton G, Fuller BJ, Seifalian AM. Vitreous cryopreservation maintains the viscoelastic property of human vascular grafts FASEB J 2006;20:874-881.[Abstract/Free Full Text]
  28. König K. Multiphoton microscopy in life sciences J Microsc 2000;200(pt 2):83-104.[Medline]
  29. Zoumi A, Yeh A, Tromberg BJ. Imaging cells and extracellular matrix in vivo by using second-harmonic generation and two-photon excited fluorescence Proc Natl Acad Sci U S A 2002;99:11014-11019.[Abstract/Free Full Text]
  30. Trask BC, Trask TM, Broekelmann T, Mecham RP. The microfibrillar proteins MAGP-1 and fibrillin-1 form a ternary complex with the chondroitin sulfate proteoglycan decorin Mol Biol Cell 2000;11:1499-1507.[Abstract/Free Full Text]
  31. Koolbergen DR, Hazekamp MG, de Heer E, et al. Structural degeneration of pulmonary homografts used as aortic valve substitute underlines early graft failure Eur J Cardiothorac Surg 2002;22:802-807.[Abstract/Free Full Text]
  32. Simionescu DT. Prevention of calcification in bioprosthetic heart valves: challenges and perspectives Expert Opin Biol Ther 2004;4:1971-1985.[Medline]
  33. Basalyga DM, Simionescu DT, Xiong W, Baxter T, Starcher BC, Vyavahare NR. Elastin degradation and calcification in an abdominal aorta injury model Circulation 2004;110:3480-3487.[Abstract/Free Full Text]
  34. Bailey M, Pillarisetti S, Jones P, Xiao H, Simionescu D, Vyavahare N. Involvement of matrix metalloproteinases and tenascin-C in elastin calcification Cardiovasc Pathol 2004;13:146-155.[Medline]
  35. Baskett RJ, Ross DB, Nanton MA, Murphy DA. Factors in the early failure of cryopreserved homograft pulmonary valves in children: preserved immunogenicity? J Thorac Cardiovasc Surg 1996;112:1170-1178.[Abstract/Free Full Text]



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K. Schenke-Layland, U. A. Stock, A. Nsair, J. Xie, E. Angelis, C. G. Fonseca, R. Larbig, A. Mahajan, K. Shivkumar, M. C. Fishbein, et al.
Cardiomyopathy is associated with structural remodelling of heart valve extracellular matrix
Eur. Heart J., June 27, 2009; (2009) ehp267v1.
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