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Virtual Journal for Biomedical Optics

Virtual Journal for Biomedical Optics

| EXPLORING THE INTERFACE OF LIGHT AND BIOMEDICINE

  • Editors: Andrew Dunn and Anthony Durkin
  • Vol. 6, Iss. 9 — Oct. 3, 2011
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Two-photon induced collagen cross-linking in bioartificial cardiac tissue

Kai Kuetemeyer, George Kensah, Marko Heidrich, Heiko Meyer, Ulrich Martin, Ina Gruh, and Alexander Heisterkamp  »View Author Affiliations


Optics Express, Vol. 19, Issue 17, pp. 15996-16007 (2011)
http://dx.doi.org/10.1364/OE.19.015996


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Abstract

Cardiac tissue engineering is a promising strategy for regenerative therapies to overcome the shortage of donor organs for transplantation. Besides contractile function, the stiffness of tissue engineered constructs is crucial to generate transplantable tissue surrogates with sufficient mechanical stability to withstand the high pressure present in the heart. Although several collagen cross-linking techniques have proven to be efficient in stabilizing biomaterials, they cannot be applied to cardiac tissue engineering, as cell death occurs in the treated area. Here, we present a novel method using femtosecond (fs) laser pulses to increase the stiffness of collagen-based tissue constructs without impairing cell viability. Raster scanning of the fs laser beam over riboflavin-treated tissue induced collagen cross-linking by two-photon photosensitized singlet oxygen production. One day post-irradiation, stress-strain measurements revealed increased tissue stiffness by around 40% being dependent on the fibroblast content in the tissue. At the same time, cells remained viable and fully functional as demonstrated by fluorescence imaging of cardiomyocyte mitochondrial activity and preservation of active contraction force. Our results indicate that two-photon induced collagen cross-linking has great potential for studying and improving artificially engineered tissue for regenerative therapies.

© 2011 OSA

1. Introduction

Tissue engineering is a promising strategy for regenerative therapies to overcome the shortage of donor organs and tissues for transplantation purposes [1

1. WHO (World Health Organization), “Cardiovascular Diseases,” Fact Sheet Number 317, Geneva, Switzerland, January2011.

]. Three-dimensional tissue engineering aims at mimicking the characteristics of natural tissue in matrix composition and morphology, as well as cellular distribution and orientation. For cardiac tissue engineering, the following physiological properties are essential for the generation of functional transplantable tissue surrogates: i) spontaneous and synchronous contractility, ii) development of sufficient systolic (contraction) and diastolic (passive) forces to support the heart’s pumping function and to withstand the hydrodynamic pressures in the heart, respectively. Several groups have shown the possibility to generate functional artificial cardiac tissue and its transplantation in animal models with beneficial effects to heart function after acute myocardial infarction [2

2. W. H. Zimmermann, C. Fink, D. Kralisch, U. Remmers, J. Weil, and T. Eschenhagen, “Three-dimensional engineered heart tissue from neonatal rat cardiac myocytes,” Biotechnol. Bioeng. 68, 106–114 (2000). [CrossRef] [PubMed]

4

4. K. L. Kreutziger and C. E. Murry, “Engineered human cardiac tissue,” Pediatr. Cardiol. 32, 334–341 (2011). [CrossRef] [PubMed]

].

Chemical cross-linking has been established as an efficient method to stabilize collagen-based biomaterials, such as porcine heart valves [12

12. C. L. McIntosh, L. L. Michaelis, A. G. Morrow, S. B. Itscoitz, D. R. Redwood, and S. E. Epstein, “Atrioventricular valve replacement with the Hancock porcine xenograft: a five-year clinical experience,” Surgery 78, 768–775 (1975). [PubMed]

] and blood vessels [13

13. H. Dardik, I. M. Ibrahim, R. Baier, S. Sprayregen, M. Levy, and I. I. Dardik, “Human umbilical cord. A new source for vascular prosthesis,” JAMA J. Am. Med. Assoc. 236, 2859–2862 (1976). [CrossRef] [CrossRef]

]. In ophthalmology, it is used to increase the human corneal stiffness to stop the progression of keratoconus [14

14. G. Wollensak, E. Spoerl, and T. Seiler, “Riboflavin/ultraviolet-A-induced collagen crosslinking for the treatment of keratoconus,” Am. J. Ophthalmol. 135, 620–627 (2003). [CrossRef] [PubMed]

]. Collagen cross-linking is generally induced by either aldehyde-reactions or UV-A irradiation with photosensitizers, such as riboflavin. Aldehyde cross-linking is presumed to result from covalent bond formation of aldehyde groups with amino groups or peptides. To avoid inflammatory reactions, cytotoxicity and calcification after implantation of cross-linked tissue into the patient, thorough removal or inactivation of excess aldehyde molecules is necessary [15

15. A. Jayakrishnan and S. R. Jameela, “Glutaraldehyde as a fixative in bioprostheses and drug delivery matrices,” Biomaterials 17, 471–484 (1996). [CrossRef] [PubMed]

]. UV-A irradiation of photosensitizers catalyzes carbonyl-based cross-linking reactions via one-photon photosensitized singlet oxygen production [16

16. M. C. DeRosa and R. J. Crutchley, “Photosensitized singlet oxygen and its applications,” Coordin. Chem. Rev. 233–234, 351–371 (2002). [CrossRef]

, 17

17. A. S. McCall, S. Kraft, H. F. Edelhauser, G. W. Kidder, R. R. Lundquist, H. E. Bradshaw, Z. Dedeic, M. J. C. Dionne, E. M. Clement, and G. W Conrad, “Mechanisms of corneal tissue cross-linking in response to treatment with topical riboflavin and long-wavelength ultraviolet radiation (UVA),” Invest. Ophthalmol. Visual Sci. 51, 129–138 (2010). [CrossRef]

]. However, the positive effect of both methods on the tissue stiffness is accompanied by cell apoptosis in the treated area. In cornea, this negative side effect is compensated by subsequent repopulation of treated areas with viable cells in vivo [18

18. G. Wollensak, E. Spoerl, M. Wilsch, and T. Seiler, “Keratocyte apoptosis after corneal collagen cross-linking using riboflavin/UVA treatment,” Cornea 23, 43–49 (2004). [CrossRef] [PubMed]

]. In case of cardiac tissue, repopulation is impaired by the non-proliferative phenotype of cardiomyocytes and most probably would result in the formation of a fibrous scar. Therefore, to apply cross-linking to cardiac tissue engineering, novel methods must be developed and evaluated which do not alter cell viability.

Compared to continuous UV-A illumination, the interaction of femtosecond (fs) laser pulses with biological tissue is based on nonlinear excitation. This enables higher penetration depths and impedes out-of-focus absorption and photodamage [25

25. K. Koenig, “Multiphoton microscopy in life sciences,” J. Microsc. 200, 83–104 (2000). [CrossRef]

]. Therefore, fs laser in the near infrared (NIR) wavelength range are extensively used for therapeutic applications on a sub-cellular level, such as chromophore assisted laser inactivation of proteins [19

19. T. Tanabe, M. Oyamada, K. Fujita, P. Dai, H. Tanaka, and T. Takamatsu, “Multiphoton excitationevoked chromophore assisted laser inactivation using green fluorescent protein,” Nat. Methods 2, 503–505 (2005). [CrossRef] [PubMed]

], intracellular nanodissection [20

20. K. Koenig, I. Riemann, P. Fischer, and K. H. Halbhuber, “Intracellular nanosurgery with near infrared femtosecond laser pulses,” Cell Mol. Biol. (Paris) 45, 195–201 (1999).

] and laser uncaging [21

21. W. Denk, J. H. Strickler, and W. W. Webb, “Two-photon laser scanning fluorescence microscopy,” Science 248, 73–76 (1990). [CrossRef] [PubMed]

]. All these applications have in common that biological samples are treated with a light-sensitive probe (e.g. fluorophore, caged compound) with a high multiphoton absorption cross-section prior to irradiation. These photosensitizers enhance the yield of chemical reactions with surrounding molecules leading to bond cleavage or rearrangements [22

22. D. Warther, S. Gug, A. Specht, F. Bolze, J. F. Nicoud, A. Mourot, and M. Goeldner, “Two-photon uncaging: new prospects in neuroscience and cellular biology,” Bioorgan. Med. Chem. 18, 7753–7758 (2010). [CrossRef]

,23

23. K. Kuetemeyer, R. Rezgui, H. Lubatschowski, and A. Heisterkamp, “Influence of laser parameters and staining on femtosecond laser-based intracellular nanosurgery,” Biomed. Opt. Express 1, 587–597 (2010). [CrossRef]

]. For example, Frederiksen et al. showed that singlet oxygen is largely produced by two-photon excitation of different photosensitizers [24

24. P. K. Frederiksen, M. Jorgensen, and P. R. Ogilby, “Two-photon photosensitized production of singlet oxygen,” J. Am. Chem. Soc. 123, 1215–1221 (2001). [CrossRef] [PubMed]

]. Great care has to be taken in the photosensitizer selection, as two-photon absorption of intrinsic fluorophores induces severe photodamage including impaired cell division or apoptosis [25

25. K. Koenig, “Multiphoton microscopy in life sciences,” J. Microsc. 200, 83–104 (2000). [CrossRef]

, 26

26. A. Hopt and E. Neher, “Highly nonlinear photodamage in two-photon fluorescence microscopy,” Biophys. J. 80, 2029–2036 (2001). [CrossRef] [PubMed]

]. At even higher laser intensities, higher-order effects become more important causing photobleaching of photosensitizers, oxidative stress and the formation of a low-density plasma [27

27. S. Kalies, K. Kuetemeyer, and A. Heisterkamp, “Mechanisms of high-order photobleaching and its relationship to intracellular ablation,” Biomed. Opt. Express 2, 805–816 (2011). [CrossRef] [PubMed]

, 28

28. A. Vogel, J. Noack, G. Huettman, and G. Paltauf, “Mechanisms of femtosecond laser nanosurgery of cells and tissues,” Appl. Phys. B 81, 1015–1047 (2005). [CrossRef]

]. As the two-photon absorption cross-section scales super-linearly with the one-photon absorption cross-section, differences between extrinsic and intrinsic fluorophores are much more pronounced in the former case [29

29. G. A. Blab, P. H. M. Lommerse, L. Cognet, G. S. Harms, and T. Schmidt, “Two-photon exciation action cross-sections of the autofluorescent proteins,” Chem. Phys. Lett. 350, 71–77 (2001). [CrossRef]

]. Consequently, nonlinear excitation with NIR wavelengths may be suitable for collagen cross-linking without compromising cell viability.

In this paper, we show the great potential of fs laser pulses for collagen cross-linking in bioartificial cardiac tissue. Collagen cross-linking was induced by raster-scanning the laser beam over riboflavin treated tissue. Subsequent stress-strain measurements were done to evaluate tissue stiffness and contractility while fluorescence microscopy was used to assess cell viability.

2. Materials and methods

2.1. Tissue preparation

Two cell sources were used for artificial tissue preparation: i) for proof-of-concept, murine embryonic fibroblasts (MEF) were used to generate a model system of collagen-rich artificial tissue, ii) neonatal rat cardiomyocytes (NRCM) were used to generate bioartificial cardiac tissue (BCT). Three-dimensional tissue was prepared as described earlier [9

9. G. Kensah, I. Gruh, J. Viering, H. Schumann, J. Dahlmann, H. Meyer, D. Skvorc, A. Baer, P. Akhyari, A. Heisterkamp, A. Haverich, and U. Martin, “A novel miniaturized multimodal bioreactor for continuous in situ assessment of bioartificial cardiac tissue during stimulation and maturation,” Tissue Eng. Pt. C Methods 17, 463–473 (2011). [CrossRef]

]. In brief, either 6·105 gamma-irradiated MEFs from CD1-ICR mice, or 1·106 freshly isolated cardiomyocytes (NRCMs) from Sprague-Dawley neonatal rats (enriched by discontinuous Percoll gradient centrifugation) were mixed with liquid extracellular matrix (consisting of 0.9 mg/ml collagen type I (R&D Systems), 10% Matrigel (BD Biosciences)) and poured into custom-made Teflon molds (220 μl). The mixture was covered with 5 ml Dulbecco’s modified Eagle’s medium (DMEM) containing 12% horse serum, 2 mM L-glutamin (all Gibco), 2% chicken embryo extract (US Biological), 10 μg/ml insulin (Sigma), 100 U/ml penicillin and 100 μg/ml streptomycin (PAA Laboratories) and cultured in a standard incubator at 5% CO2, 37°C and 80% humidity with daily medium exchange. Within the molds, the solidified tissue was suspended between two titanium rods and had an average cross-sectional area of 0.8 mm2 on day 7 (see Fig. 1a).

Fig. 1 (a) Phase contrast microscopy image of a MEF-based tissue suspended between titanium rods on day 7 prior to laser irradiation. Scale bar: 1 mm. (b) Schematic setup for two-photon induced collagen cross-linking. L1 / L2: f1=250 and f2=−50 mm planoconvex and plano-concave lenses; L3: focusing lens with f=140 mm or f=400 mm. (c) Tissue constructs were raster-scanned with a constant scanning speed and line separation Δy being equal to the focal spot radius.

All experiments were performed in accordance with the principles of Laboratory Animal Care (NIH publication No. 86-23, revised 1985) as well as the Animal Welfare Law of Lower Saxony, Germany.

2.2. Optical Setup

The optical setup is shown in Fig. 1b. The laser source was a regeneratively amplified Ti:Sapphire laser system producing femtosecond pulses, either a Spectra Physics Spitfire Pro (120 fs, 800 nm, 5 kHz) or a Thales Bright (120 fs, 780 nm, 5 kHz). At the used wavelengths, riboflavin has a two-photon action cross-section of about 0.5 GM at 780 nm and 0.45 GM at 800 nm [30

30. W. R. Zipfel, R. M. Williams, R. Christie, A. Y. Nikitin, B. T. Hyman, and W. W. Webb, “Live tissue intrinsic emission microscopy using multiphoton-excited native fluorescence and second harmonic generation,” Proc. Natl. Acad. Sci. U.S.A. 100, 7075–7080 (2003). [CrossRef] [PubMed]

]. The laser beam was demagnified by a two-lens telescope to match the diameter of the following optics and attenuated by a variable neutral density (ND) filter. Two high-speed galvanometer mirrors (Litrack) were used to scan the laser beam in the x-y plane. Focusing into the tissue was achieved either by a 140-mm or 400-mm focal length lens. This resulted in a spot diameter of 130 and 380 μm as well as a rayleigh length of 17 and 145 mm, respectively, being much larger than the average tissue depth of 0.8 mm.

2.3. Two-photon induced collagen cross-linking

After six days of cultivation, the photosensitizer riboflavin (RF, Sigma Aldrich) was added to one part of the tissue constructs at 0.01% (0.27 mM) concentration in the culture medium. This RF concentration is one order below that used for UV-A collagen cross-linking [14

14. G. Wollensak, E. Spoerl, and T. Seiler, “Riboflavin/ultraviolet-A-induced collagen crosslinking for the treatment of keratoconus,” Am. J. Ophthalmol. 135, 620–627 (2003). [CrossRef] [PubMed]

]. However, solutions with higher concentrations could not be made because of its limited solubility in the culture medium. On day seven, each tissue was transferred within the Teflon mold to a 35 mm glass bottom dish (ibidi GmbH) with a thickness of 170 μm. Collagen cross-linking was induced by raster scanning the laser beam over the tissue with a constant scanning speed and line separation (see Fig. 1c). The line separation Δy corresponded to the focal spot radius for both focusing lenses. Depending on the scanning parameters, the cross-linking procedure lasted approximately 30 seconds to 5 min. The experiments were done at room temperature and normal atmosphere unless otherwise stated. Immediately after laser irradiation, each tissue was washed in culture medium without RF and further incubated for 24 hours at 37°C and 5% CO2 humidified atmosphere.

2.4. Stress-strain measurements

Forces were measured in a bioreactor system that allows for continuous increase in strain and on-line force measurement in a standard incubator [9

9. G. Kensah, I. Gruh, J. Viering, H. Schumann, J. Dahlmann, H. Meyer, D. Skvorc, A. Baer, P. Akhyari, A. Heisterkamp, A. Haverich, and U. Martin, “A novel miniaturized multimodal bioreactor for continuous in situ assessment of bioartificial cardiac tissue during stimulation and maturation,” Tissue Eng. Pt. C Methods 17, 463–473 (2011). [CrossRef]

]. For MEF-based and bioartificial cardiac tissue (BCT), the increase in passive force as a function of increase in strain (33.25 ± 1.75 μm/s) along the longitudinal axis of the tissue constructs was measured for each tissue in culture medium until the preload reached 20%. The engineering stress was calculated as force divided by the initial cross-sectional area. The slope of the linear region of the stress-strain curve (”Young’s modulus”) was determined by linear regression. To assess the active contractility of treated and untreated control groups of cardiac tissue, BCTs were electrically paced five times with no preload at 25 V with rectangular pulses (5 ms) prior to stress-strain measurements.

The statistical significance was tested using one-way analysis of variance (ANOVA) followed by multiple comparisons against the untreated control group using Dunnett’s test. Differences were considered significant at P < 0.05.

2.5. Fluorescence microscopy

One day prior to cross-linking experiments, the tissue was incubated with 25 nM tetramethylrhodamine methyl ester (TMRM; Invitrogen). TMRM is a cationic fluorescent dye being rapidly and reversibly taken up by live cells and sequestered to active mitochondria, and therefore can be used to monitor the cellular metabolic activity. Re-staining with TMRM immediately after laser irradiation was performed to compensate for photobleaching. Images of the tissue cross-sectional area were obtained while sandwiched between two cover glasses. To analyze the volumetric cell density one day after cross-linking, tissues were fixed for 20 minutes in acetone and nuclei were stained with DAPI (Invitrogen). Fluorescence images were either captured with a commercial AxioObserver Z1 microscope (Carl Zeiss AG) or a custom made scanning laser optical tomograph (SLOT) described elsewhere [31

31. R. A. Lorbeer, M. Heidrich, C. Lorbeer, D. F. Ramirez-Ojeda, G. Bicker, H. Meyer, and A. Heisterkamp, “Highly efficient 3D fluorescence microscopy with a scanning laser optical tomograph,” Opt. Express 19, 5419–5430 (2011). [CrossRef] [PubMed]

]. In brief, SLOT is a highly efficient 3D fluorescence microscopy technique capable of imaging specimens with sizes up to several millimeters. For imaging, the specimen was mounted in a glass capillary filled with 100% glycerol for optical clearing.

3. Results

3.1. Collagen cross-linking in MEF-based tissue

The first set of experiments was done with the Spitfire Pro laser system. The focal spot radius and scanning speed were determined to 65 μm and 300 μm/s, respectively, corresponding to an irradiation time of approximately 5 minutes. In preliminary experiments, tissues were irradiated with different pulse energies to identify suitable parameters. The laser fluence was thereby defined as the pulse energy, divided by the focal area, times the number of pulses. Two laser fluences were chosen empirically for two-photon induced collagen cross-linking [28

28. A. Vogel, J. Noack, G. Huettman, and G. Paltauf, “Mechanisms of femtosecond laser nanosurgery of cells and tissues,” Appl. Phys. B 81, 1015–1047 (2005). [CrossRef]

]: 160 and 320 J/cm2. For MEF-based tissue as our model system of collagen-rich artificial tissue, samples were divided into five groups: (1) untreated control, (2) RF treatment, (3) 160 J/cm2 laser fluence with RF treatment and 320 J/cm2 laser fluence (4) with and (5) without RF treatment.

All tissues exhibited similar stress-strain relationships 24 hours after irradiation. The stress-strain curves showed an initial nonlinear (”toe”) region followed by a linear region from about 15% strain (see Fig. 2a). RF treatment alone did not influence the stress-strain relation. By contrast, irradiation of RF treated tissues at 160 J/cm2 resulted in an increased stiffness of 35% compared to untreated controls at 20% strain (see Fig. 2b). A similar rise was observed for the slope of the stress-strain relation in the linear region, the so-called Young’s modulus (46.6±0.3 vs. 34.8±2.3 kPa). When the laser fluence was doubled to 320 J/cm2, the positive effect on tissue stiffness was no longer observed, independent of RF treatment.

Fig. 2 (a) Stress-strain relation of MEF-based tissues after RF treatment and fs laser irradiation. RF or laser treatment alone did not have an effect on tissue stiffness. Increased stiffening occurred after RF treatment and irradiation at 160 J/cm2. At the higher fluence of 320 J/cm2, this positive effect was no longer observed. (b) Irradiation of RF treated tissues at 160 J/cm2 resulted in an increase of engineering stress at 20% strain and Young’s modulus by 35% compared to untreated controls. Each data point represents the mean ± SEM of two experiments.

At the lower fluence, the TMRM and DAPI fluorescence intensities of irradiated and non-irradiated areas were comparable. Therefore, laser irradiation at these parameters had no detrimental effects on cell metabolic activity and density. By contrast, the metabolic activity was markedly reduced at the higher fluence (see Fig. 4a), independent of RF treatment. At the same time, no influence on cell density was found as indicated by unchanged DAPI fluorescence intensities (data not shown).

Fig. 4 (a) Average TMRM fluorescence intensity in untreated control and RF treated + laser irradiated MEF-based tissues and BCTs. Each bar represents the mean ± SEM of two (MEF) and five (BCT) measurements. (b) Phase contrast and TMRM fluorescence images over the cross-sectional area of a BCT treated with RF and irradiated from below at 300 J/cm2. The maximum depth of laser treatment was measured at an incident angle of 0° to about 520 μm. Scale bar: 200 μm.

3.2. Collagen cross-linking in bioartificial cardiac tissue (BCT)

As with MEF-based tissue, the tensile stress showed a nonlinear dependence on the strain amplitude, which changes into a linear dependence at 15% strain. However, the stiffness and Young’s modulus of BCTs were about a factor three lower (data not shown). Irradiation of untreated BCTs at 160 J/cm2 did not change the stress-strain relation. Using the same irradiation parameters for RF treated tissues resulted in an increased stiffness by 25% compared to untreated controls at 20% strain, while Young’s modulus only increased by 6% (14.9±1.7 vs. 14.1±1.4 kPa). At the higher fluence of 320 J/cm2, mechanical properties of tissues were no longer influenced. Because of the high variation of stiffness in all treatment groups, the measured differences were not statistically significant (P > 0.25).

Fig. 3 Representative (a,b) TMRM and (c,d) DAPI fluorescence microscopy images of RF treated and laser irradiated BCTs: (a,c) 160 J/cm2 laser fluence + RF; (b,d) 320 J/cm2 laser fluence + RF. The white dotted lines separate the irradiated (bottom) and non-irradiated areas (top). A high magnification image of the boxed area in (e) is provided in (b). At the higher laser fluence, irradiation resulted in a marked decrease of TMRM fluorescence intensity, while DAPI fluorescence was still intense. Scale bar: 500 μm.
Fig. 5 Representative SLOT fluorescence images of cross-linked BCT after fixation and DAPI staining. The section plane of the right image is indicated by the red dashed line in the left image. No difference in the volumetric cell density was observed between irradiated and non-irradiated areas. Scale bar: 200 μm.

In case of BCTs, tissue functionality can be directly assessed by measurement of contractile forces, which will be exerted by viable cardiomyocytes only. All BCT groups exhibited spontaneous active contraction with no preload both before and 24 hours after irradiation. Electrical stimulation in the bioreactor induced contraction forces up to 0.8 mN with no significant difference between the experimental groups (P > 0.3, data not shown).

3.3. Optimization of two-photon induced collagen cross-linking in BCTs

For the treatment of cardiac tissue, protocols were further optimized to reduce the stress applied to the cells and to improve the enhancement of mechanical properties. First, an infrared lamp was integrated into the setup and HEPES buffer was added to the culture medium to maintain optimal culture conditions outside the incubator (37°C and 5% CO2). Second, the focal spot diameter and scanning speed were increased from 130 to 380 μm and from 300 to 1300 μm/s, respectively, to reduce the irradiation time to 30 s. Third, repetitive raster scanning of the tissue enabled the use of lower pulse energies for collagen cross-linking as multiphoton-induced photochemical effects accumulate over multiple pulses [23

23. K. Kuetemeyer, R. Rezgui, H. Lubatschowski, and A. Heisterkamp, “Influence of laser parameters and staining on femtosecond laser-based intracellular nanosurgery,” Biomed. Opt. Express 1, 587–597 (2010). [CrossRef]

, 28

28. A. Vogel, J. Noack, G. Huettman, and G. Paltauf, “Mechanisms of femtosecond laser nanosurgery of cells and tissues,” Appl. Phys. B 81, 1015–1047 (2005). [CrossRef]

]. However, to maintain a high sample throughput, the scanning pattern was only applied five times. Fourth, the experiments were done with the Thales Bright laser system at 780 nm at which the two-photon action cross-section of RF is about 10% higher [30

30. W. R. Zipfel, R. M. Williams, R. Christie, A. Y. Nikitin, B. T. Hyman, and W. W. Webb, “Live tissue intrinsic emission microscopy using multiphoton-excited native fluorescence and second harmonic generation,” Proc. Natl. Acad. Sci. U.S.A. 100, 7075–7080 (2003). [CrossRef] [PubMed]

].

Using optimized conditions, stress-strain measurements revealed a strong influence of the laser fluence on the mechanical properties (see Fig. 6). Significant tissue stiffening by 40% at 20% strain was only observed in a small window of laser fluences around 50 J/cm2 (P < 0.05). The corresponding Young’s modulus increased by about 15% (30.6±0.9 vs. 26.5±1.9 kPa). Compared to Sec. 3.2, the enhancement of mechanical properties was improved by about a factor two. At the same time, no significant influence on active contraction force and cell metabolic activity was observed (see Fig. 4a), independent of the laser fluence up to 95 J/cm2 (P > 0.1).

Fig. 6 (a) Stress-strain relation of bioartificial cardiac tissue (BCT) after optimization of the cross-linking procedure. Significant stiffening was observed in a small process window around 50 J/cm2. (b) Laser irradiation of RF treated BCTs at 50 J/cm2 resulted in a significantly increased stiffness by 40% at 20 % strain, while the active contraction force was comparable to untreated controls. Each data point represents the mean ± SEM of at least four experiments. * P<0.05 versus untreated control group.

4. Discussion and conclusion

The presented results indicate the great potential of femtosecond (fs) laser pulses for fast, efficient and non-toxic cross-linking of bioartificial collagenous tissue.

To induce collagen cross-linking, we irradiated riboflavin treated collagenous tissue with fs laser pulses in the NIR wavelength range at low pulse energies generally used for multiphoton-induced photochemistry [28

28. A. Vogel, J. Noack, G. Huettman, and G. Paltauf, “Mechanisms of femtosecond laser nanosurgery of cells and tissues,” Appl. Phys. B 81, 1015–1047 (2005). [CrossRef]

]. The resulting maximum treatment depth was about 520 μm, corresponding to 65% of the average tissue depth (see Fig. 4b). In contrast to existing methods, raster scanning of the laser beam enabled full spatial control of selective cross-linking in three-dimensional tissue [14

14. G. Wollensak, E. Spoerl, and T. Seiler, “Riboflavin/ultraviolet-A-induced collagen crosslinking for the treatment of keratoconus,” Am. J. Ophthalmol. 135, 620–627 (2003). [CrossRef] [PubMed]

, 15

15. A. Jayakrishnan and S. R. Jameela, “Glutaraldehyde as a fixative in bioprostheses and drug delivery matrices,” Biomaterials 17, 471–484 (1996). [CrossRef] [PubMed]

]. Owing to the surface curvature, the coupling efficiency of laser energy into tissue continuously decreased towards the edges of the tissue, as described by the Fresnel reflectivity equations and Snell’s Law [32

32. M. H. Niemz, Laser-Tissue Interactions: Fundamentals and Applications (Springer, 2007).

]. Consequently, only half the tissue volume was irradiated with about the same fluence. The achievable precision with our setup was in the order of 100 μm, equivalent to the focal spot diameter. By applying tight focusing with high numerical aperture objectives, the focal spot diameter can be reduced to the sub-micrometer range [25

25. K. Koenig, “Multiphoton microscopy in life sciences,” J. Microsc. 200, 83–104 (2000). [CrossRef]

]. In combination with a rotating stage for the tissue, homogeneous energy deposition and hence selective cross-linking within a sub-femtoliter volume is feasible in future experiments.

In our experiments, we used MEF-based and bioartificial cardiac tissue (BCT), containing fibroblasts and enriched primary cardiomyocytes with a lower proportion of fibroblasts, respectively, to examine the influence of different cell compositions. One-day post irradiation, the stress-strain relations of both tissue types exhibited a nonlinear ”toe” region followed by a linear region from about 15% strain, characteristic of collagenous tissue [33

33. B. A. Roeder, K. Kokini, J. E. Sturgis, J. P. Robinson, and S. L. Voytik-Harbin, “Tensile mechanical properties of three-dimensional type I collagen extracellular matrices with varied microstructure,” J. Biomech. Eng. 124, 214–222 (2002). [CrossRef] [PubMed]

, 34

34. G. Wollensak, E. Spoerl, and T. Seiler, “Stress-strain measurements of human and porcine corneas after riboflavinultraviolet-A-induced cross-linking,” J. Cataract Refractive Surg. 29, 1780–1785 (2003). [CrossRef]

]. At a fluence of 160 J/cm2, considerable tissue stiffening and increase in Young’s modulus by 35% was observed in MEF-based tissues. Using the same parameters, the enhancement of mechanical properties was lower in BCTs (compare Figs. 2a and 6a). The higher increase in MEF-based tissues can be explained by the higher initial fibroblast content and hence a higher content and/or turnover of extracellular matrix (ECM) molecules, such as collagen, over cultivation time [35

35. M. Eghbali and K. T. Weber, “Collagen and the myocardium: fibrillar structure, biosynthesis and degradation in relation to hypertrophy and its regression,” Mol. Cell. Biochem. 96, 1–14 (1990). [CrossRef] [PubMed]

].

As riboflavin is known to induce UV-A collagen cross-linking via one-photon photosensitized singlet oxygen production [16

16. M. C. DeRosa and R. J. Crutchley, “Photosensitized singlet oxygen and its applications,” Coordin. Chem. Rev. 233–234, 351–371 (2002). [CrossRef]

, 17

17. A. S. McCall, S. Kraft, H. F. Edelhauser, G. W. Kidder, R. R. Lundquist, H. E. Bradshaw, Z. Dedeic, M. J. C. Dionne, E. M. Clement, and G. W Conrad, “Mechanisms of corneal tissue cross-linking in response to treatment with topical riboflavin and long-wavelength ultraviolet radiation (UVA),” Invest. Ophthalmol. Visual Sci. 51, 129–138 (2010). [CrossRef]

], the observed increase in stiffness most likely resulted from similar processes upon two-photon excitation. This is underlined by two facts: i) riboflavin fluorescence was observed during fs laser beam raster scanning and ii) irradiation of untreated tissues did not influence the tissue stiffness. Although TMRM was present in the culture medium with a much higher two-photon action cross-section than riboflavin [36

36. C. Xu and W. W. Webb “Measurement of two-photon excitation cross sections of molecular fluorophores with data from 690 to 1050 nm,” J. Opt. Soc. Am. B 13, 481–491 (1996). [CrossRef]

], it had no detectable influence on collagen cross-linking owing to the 10,000 times lower concentration and the selective accumulation in mitochondria.

UV-A cross-linking protocols for corneas are usually associated with cell death within the treated area followed by in-vivo repopulation [18

18. G. Wollensak, E. Spoerl, M. Wilsch, and T. Seiler, “Keratocyte apoptosis after corneal collagen cross-linking using riboflavin/UVA treatment,” Cornea 23, 43–49 (2004). [CrossRef] [PubMed]

]. In our approach, cell viability was maintained both in MEF-based tissue and BCT, as indicated by strong TMRM fluorescence in the irradiated area (see Figs. 3a and 4a). Furthermore, their volumetric density was not influenced and DAPI fluorescence was still intense (see Figs. 5 and 3c). As active contraction forces of irradiated BCTs were comparable to untreated controls (see Fig. 6b), we can conclude that cardiomyocytes remained fully functional after cross-linking. This could indicate that intracellular free radical scavengers like glutathione were able to neutralize the oxidative stress induced by fs laser irradiation [25

25. K. Koenig, “Multiphoton microscopy in life sciences,” J. Microsc. 200, 83–104 (2000). [CrossRef]

, 37

37. B. P. Yu, “Cellular defenses against damage from reactive oxygen species,” Physiol. Rev. 74, 139–162 (1994). [PubMed]

]. To the best of our knowledge, similar observations have only been made after one-photon induced cross-linking of fibrin-based engineered connective tissue [38

38. Z. H. Syedain, J. Bjork, L. Sando, and R. T. Tranquillo, “Controlled compaction with ruthenium-catalyzed photochemical cross-linking of fibrin-based engineered connective tissue,” Biomaterials 30, 6695–6701 (2009). [CrossRef] [PubMed]

]. Therefore, our experimental setup provides for the first time a fast and effective method to increase the stiffness of collagenous tissue without impairing cell viability. However, great care has to be taken in adjusting the laser fluence, as cellular metabolic activity markedly decreased and no positive effect on tissue stiffness was observed at very high fluences (see Fig. 3b and 2a). This is likely a result of excessive oxidative stress and/or progressive heat accumulation within the tissue during the scanning procedure [39

39. A. Vogel and V. Venugopalan, “Mechanisms of pulsed laser ablation of biological tissues,” Chem. Rev. 103, 577–644 (2003). [CrossRef] [PubMed]

]. Nevertheless, active contraction forces of those BCTs were comparable to untreated controls (see Fig. 6b)), providing evidence that cardiomyocytes were still functional.

Our results suggest a ”window” of laser fluences, in which two-photon induced collagen cross-linking can be achieved while maintaining cell viability. To the best of our knowledge, this was not yet observed using one-photon absorption. To obtain a safe process window, the difference of the absorption cross-sections of riboflavin compared to intrinsic fluorophores, such as NAD(P)H, has to be sufficiently high [25

25. K. Koenig, “Multiphoton microscopy in life sciences,” J. Microsc. 200, 83–104 (2000). [CrossRef]

]. Other groups have shown that the two-photon absorption cross-section scales super-linearly with the one-photon absorption cross-section [29

29. G. A. Blab, P. H. M. Lommerse, L. Cognet, G. S. Harms, and T. Schmidt, “Two-photon exciation action cross-sections of the autofluorescent proteins,” Chem. Phys. Lett. 350, 71–77 (2001). [CrossRef]

]. Therefore, we assume that nonlinear excitation of riboflavin with NIR wavelengths is the only mechanism providing a large enough process window for minimally invasive collagen cross-linking. Future studies have to further optimize the laser fluence within the process window to enhance the achievable increase in stiffness.

In conclusion, we demonstrated for the first time the fast and efficient cross-linking of riboflavin treated collagenous tissues with fs laser pulses in the NIR wavelength range. In contrast to existing methods, the laser-tissue interaction is based on nonlinear (two-photon) absorption. Our experimental data suggest that this provides a process window, in which collagen cross-linking is achieved while maintaining cell viability and functionality. The nonlinear laser-tissue interaction in combination with high spatial precision in the micrometer range offers the possibility for selective cross-linking of arbitrarily defined patterns in three-dimensional tissue engineered constructs.

Patterning of artificial cardiac tissue could be implemented to recapitulate the intrinsic anisotropy of native myocardial tissue. As tissue stiffness plays an important role in remodeling and regeneration after myocardial infarction, artificial tissue with precisely defined mechanical properties using two-photon induced cross-linking could provide new therapeutic strategies to mechanically support heart function. For instance, it was proposed that most of the benefit could be achieved by selectively stiffening the longitudinal direction without altering the circumferential direction [40

40. G. M. Fomovsky, J. R. Macadangdang, G. Ailawadi, and J. W. Holmes, “Model-based design of mechanical therapies for myocardial infarction,” J. Cardiovasc. Transl. Res. 4, 82–91 (2011). [CrossRef]

]. Therefore, our fast, efficient and cell-compatible method for controlling the physical properties of the ECM is a valuable tool for studying and improving engineered cardiac tissue for regenerative therapies.

Acknowledgments

We would like to thank David Skvorc for excellent technical assistance. This work is supported by funding from the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) within the Cluster of Excellence ”REBIRTH” (From Regenerative Biology to Reconstructive Therapy).

References and links

1.

WHO (World Health Organization), “Cardiovascular Diseases,” Fact Sheet Number 317, Geneva, Switzerland, January2011.

2.

W. H. Zimmermann, C. Fink, D. Kralisch, U. Remmers, J. Weil, and T. Eschenhagen, “Three-dimensional engineered heart tissue from neonatal rat cardiac myocytes,” Biotechnol. Bioeng. 68, 106–114 (2000). [CrossRef] [PubMed]

3.

W. H. Zimmermann, M. Tiburcy, and T. Eschenhagen, “Cardiac tissue engineering: a clinical perspective,” Future Cardiol. 3, 435–445 (2007). [CrossRef] [PubMed]

4.

K. L. Kreutziger and C. E. Murry, “Engineered human cardiac tissue,” Pediatr. Cardiol. 32, 334–341 (2011). [CrossRef] [PubMed]

5.

B. Bhana, R. K. Iyer, W. L. Chen, R. Zhao, K. L. Sider, M. Likhitpanichkul, C. A. Simmons, and M. Radisic, “Influence of substrate stiffness on the phenotype of heart cells,” Biotechnol. Bioeng. 105, 1148–1160 (2010).

6.

L. Moeller, A. Krause, J. Dahlmann, I. Gruh, A. Kirschning, and G. Draeger, “Preparation and evaluation of hydrogel-composites from methacrylated hyaluronic acid, alginate, and gelatin for tissue engineering,” Int. J. Artif. Organs 34, 93–102 (2011). [CrossRef]

7.

A. Marsano, R. Maidhof, L. Q. Wan, Y. Wang, J. Gao, N. Tandon, and G. Vunjak-Novakovic, “Scaffold stiffness affects the contractile function of three-dimensional engineered cardiac constructs,” Biotechnol. Prog. 26, 1382–1390 (2010). [CrossRef] [PubMed]

8.

C. Fink, S. Ergun, D. Kralisch, U. Remmers, J. Weil, and T. Eschenhagen, “Chronic stretch of engineered heart tissue induces hypertrophy and functional improvement,” FASEB J. 14, 669–679 (2000). [PubMed]

9.

G. Kensah, I. Gruh, J. Viering, H. Schumann, J. Dahlmann, H. Meyer, D. Skvorc, A. Baer, P. Akhyari, A. Heisterkamp, A. Haverich, and U. Martin, “A novel miniaturized multimodal bioreactor for continuous in situ assessment of bioartificial cardiac tissue during stimulation and maturation,” Tissue Eng. Pt. C Methods 17, 463–473 (2011). [CrossRef]

10.

W. M. Elbjeirami, E. O. Yonter, B. C. Starcher, and J. L. West, “Enhancing mechanical properties of tissue-engineered constructs via lysyl oxidase crosslinking activity,” J. Biomed. Mater. Res. 66, 513–521 (2003). [CrossRef]

11.

T. S. Girton, T. R. Oegema, and R. T. Tranquillo, “Exploiting glycation to stiffen and strengthen tissue equivalents for tissue engineering,” J. Biomed. Mater. Res. 46, 87–92 (1999). [CrossRef] [PubMed]

12.

C. L. McIntosh, L. L. Michaelis, A. G. Morrow, S. B. Itscoitz, D. R. Redwood, and S. E. Epstein, “Atrioventricular valve replacement with the Hancock porcine xenograft: a five-year clinical experience,” Surgery 78, 768–775 (1975). [PubMed]

13.

H. Dardik, I. M. Ibrahim, R. Baier, S. Sprayregen, M. Levy, and I. I. Dardik, “Human umbilical cord. A new source for vascular prosthesis,” JAMA J. Am. Med. Assoc. 236, 2859–2862 (1976). [CrossRef] [CrossRef]

14.

G. Wollensak, E. Spoerl, and T. Seiler, “Riboflavin/ultraviolet-A-induced collagen crosslinking for the treatment of keratoconus,” Am. J. Ophthalmol. 135, 620–627 (2003). [CrossRef] [PubMed]

15.

A. Jayakrishnan and S. R. Jameela, “Glutaraldehyde as a fixative in bioprostheses and drug delivery matrices,” Biomaterials 17, 471–484 (1996). [CrossRef] [PubMed]

16.

M. C. DeRosa and R. J. Crutchley, “Photosensitized singlet oxygen and its applications,” Coordin. Chem. Rev. 233–234, 351–371 (2002). [CrossRef]

17.

A. S. McCall, S. Kraft, H. F. Edelhauser, G. W. Kidder, R. R. Lundquist, H. E. Bradshaw, Z. Dedeic, M. J. C. Dionne, E. M. Clement, and G. W Conrad, “Mechanisms of corneal tissue cross-linking in response to treatment with topical riboflavin and long-wavelength ultraviolet radiation (UVA),” Invest. Ophthalmol. Visual Sci. 51, 129–138 (2010). [CrossRef]

18.

G. Wollensak, E. Spoerl, M. Wilsch, and T. Seiler, “Keratocyte apoptosis after corneal collagen cross-linking using riboflavin/UVA treatment,” Cornea 23, 43–49 (2004). [CrossRef] [PubMed]

19.

T. Tanabe, M. Oyamada, K. Fujita, P. Dai, H. Tanaka, and T. Takamatsu, “Multiphoton excitationevoked chromophore assisted laser inactivation using green fluorescent protein,” Nat. Methods 2, 503–505 (2005). [CrossRef] [PubMed]

20.

K. Koenig, I. Riemann, P. Fischer, and K. H. Halbhuber, “Intracellular nanosurgery with near infrared femtosecond laser pulses,” Cell Mol. Biol. (Paris) 45, 195–201 (1999).

21.

W. Denk, J. H. Strickler, and W. W. Webb, “Two-photon laser scanning fluorescence microscopy,” Science 248, 73–76 (1990). [CrossRef] [PubMed]

22.

D. Warther, S. Gug, A. Specht, F. Bolze, J. F. Nicoud, A. Mourot, and M. Goeldner, “Two-photon uncaging: new prospects in neuroscience and cellular biology,” Bioorgan. Med. Chem. 18, 7753–7758 (2010). [CrossRef]

23.

K. Kuetemeyer, R. Rezgui, H. Lubatschowski, and A. Heisterkamp, “Influence of laser parameters and staining on femtosecond laser-based intracellular nanosurgery,” Biomed. Opt. Express 1, 587–597 (2010). [CrossRef]

24.

P. K. Frederiksen, M. Jorgensen, and P. R. Ogilby, “Two-photon photosensitized production of singlet oxygen,” J. Am. Chem. Soc. 123, 1215–1221 (2001). [CrossRef] [PubMed]

25.

K. Koenig, “Multiphoton microscopy in life sciences,” J. Microsc. 200, 83–104 (2000). [CrossRef]

26.

A. Hopt and E. Neher, “Highly nonlinear photodamage in two-photon fluorescence microscopy,” Biophys. J. 80, 2029–2036 (2001). [CrossRef] [PubMed]

27.

S. Kalies, K. Kuetemeyer, and A. Heisterkamp, “Mechanisms of high-order photobleaching and its relationship to intracellular ablation,” Biomed. Opt. Express 2, 805–816 (2011). [CrossRef] [PubMed]

28.

A. Vogel, J. Noack, G. Huettman, and G. Paltauf, “Mechanisms of femtosecond laser nanosurgery of cells and tissues,” Appl. Phys. B 81, 1015–1047 (2005). [CrossRef]

29.

G. A. Blab, P. H. M. Lommerse, L. Cognet, G. S. Harms, and T. Schmidt, “Two-photon exciation action cross-sections of the autofluorescent proteins,” Chem. Phys. Lett. 350, 71–77 (2001). [CrossRef]

30.

W. R. Zipfel, R. M. Williams, R. Christie, A. Y. Nikitin, B. T. Hyman, and W. W. Webb, “Live tissue intrinsic emission microscopy using multiphoton-excited native fluorescence and second harmonic generation,” Proc. Natl. Acad. Sci. U.S.A. 100, 7075–7080 (2003). [CrossRef] [PubMed]

31.

R. A. Lorbeer, M. Heidrich, C. Lorbeer, D. F. Ramirez-Ojeda, G. Bicker, H. Meyer, and A. Heisterkamp, “Highly efficient 3D fluorescence microscopy with a scanning laser optical tomograph,” Opt. Express 19, 5419–5430 (2011). [CrossRef] [PubMed]

32.

M. H. Niemz, Laser-Tissue Interactions: Fundamentals and Applications (Springer, 2007).

33.

B. A. Roeder, K. Kokini, J. E. Sturgis, J. P. Robinson, and S. L. Voytik-Harbin, “Tensile mechanical properties of three-dimensional type I collagen extracellular matrices with varied microstructure,” J. Biomech. Eng. 124, 214–222 (2002). [CrossRef] [PubMed]

34.

G. Wollensak, E. Spoerl, and T. Seiler, “Stress-strain measurements of human and porcine corneas after riboflavinultraviolet-A-induced cross-linking,” J. Cataract Refractive Surg. 29, 1780–1785 (2003). [CrossRef]

35.

M. Eghbali and K. T. Weber, “Collagen and the myocardium: fibrillar structure, biosynthesis and degradation in relation to hypertrophy and its regression,” Mol. Cell. Biochem. 96, 1–14 (1990). [CrossRef] [PubMed]

36.

C. Xu and W. W. Webb “Measurement of two-photon excitation cross sections of molecular fluorophores with data from 690 to 1050 nm,” J. Opt. Soc. Am. B 13, 481–491 (1996). [CrossRef]

37.

B. P. Yu, “Cellular defenses against damage from reactive oxygen species,” Physiol. Rev. 74, 139–162 (1994). [PubMed]

38.

Z. H. Syedain, J. Bjork, L. Sando, and R. T. Tranquillo, “Controlled compaction with ruthenium-catalyzed photochemical cross-linking of fibrin-based engineered connective tissue,” Biomaterials 30, 6695–6701 (2009). [CrossRef] [PubMed]

39.

A. Vogel and V. Venugopalan, “Mechanisms of pulsed laser ablation of biological tissues,” Chem. Rev. 103, 577–644 (2003). [CrossRef] [PubMed]

40.

G. M. Fomovsky, J. R. Macadangdang, G. Ailawadi, and J. W. Holmes, “Model-based design of mechanical therapies for myocardial infarction,” J. Cardiovasc. Transl. Res. 4, 82–91 (2011). [CrossRef]

OCIS Codes
(170.1610) Medical optics and biotechnology : Clinical applications
(190.4180) Nonlinear optics : Multiphoton processes
(260.5130) Physical optics : Photochemistry

ToC Category:
Medical Optics and Biotechnology

History
Original Manuscript: May 31, 2011
Revised Manuscript: July 15, 2011
Manuscript Accepted: July 18, 2011
Published: August 5, 2011

Virtual Issues
Vol. 6, Iss. 9 Virtual Journal for Biomedical Optics

Citation
Kai Kuetemeyer, George Kensah, Marko Heidrich, Heiko Meyer, Ulrich Martin, Ina Gruh, and Alexander Heisterkamp, "Two-photon induced collagen cross-linking in bioartificial cardiac tissue," Opt. Express 19, 15996-16007 (2011)
http://www.opticsinfobase.org/vjbo/abstract.cfm?URI=oe-19-17-15996


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References

  1. WHO (World Health Organization), “Cardiovascular Diseases,” Fact Sheet Number 317, Geneva, Switzerland, January2011.
  2. W. H. Zimmermann, C. Fink, D. Kralisch, U. Remmers, J. Weil, and T. Eschenhagen, “Three-dimensional engineered heart tissue from neonatal rat cardiac myocytes,” Biotechnol. Bioeng. 68, 106–114 (2000). [CrossRef] [PubMed]
  3. W. H. Zimmermann, M. Tiburcy, and T. Eschenhagen, “Cardiac tissue engineering: a clinical perspective,” Future Cardiol. 3, 435–445 (2007). [CrossRef] [PubMed]
  4. K. L. Kreutziger and C. E. Murry, “Engineered human cardiac tissue,” Pediatr. Cardiol. 32, 334–341 (2011). [CrossRef] [PubMed]
  5. B. Bhana, R. K. Iyer, W. L. Chen, R. Zhao, K. L. Sider, M. Likhitpanichkul, C. A. Simmons, and M. Radisic, “Influence of substrate stiffness on the phenotype of heart cells,” Biotechnol. Bioeng. 105, 1148–1160 (2010).
  6. L. Moeller, A. Krause, J. Dahlmann, I. Gruh, A. Kirschning, and G. Draeger, “Preparation and evaluation of hydrogel-composites from methacrylated hyaluronic acid, alginate, and gelatin for tissue engineering,” Int. J. Artif. Organs 34, 93–102 (2011). [CrossRef]
  7. A. Marsano, R. Maidhof, L. Q. Wan, Y. Wang, J. Gao, N. Tandon, and G. Vunjak-Novakovic, “Scaffold stiffness affects the contractile function of three-dimensional engineered cardiac constructs,” Biotechnol. Prog. 26, 1382–1390 (2010). [CrossRef] [PubMed]
  8. C. Fink, S. Ergun, D. Kralisch, U. Remmers, J. Weil, and T. Eschenhagen, “Chronic stretch of engineered heart tissue induces hypertrophy and functional improvement,” FASEB J. 14, 669–679 (2000). [PubMed]
  9. G. Kensah, I. Gruh, J. Viering, H. Schumann, J. Dahlmann, H. Meyer, D. Skvorc, A. Baer, P. Akhyari, A. Heisterkamp, A. Haverich, and U. Martin, “A novel miniaturized multimodal bioreactor for continuous in situ assessment of bioartificial cardiac tissue during stimulation and maturation,” Tissue Eng. Pt. C Methods 17, 463–473 (2011). [CrossRef]
  10. W. M. Elbjeirami, E. O. Yonter, B. C. Starcher, and J. L. West, “Enhancing mechanical properties of tissue-engineered constructs via lysyl oxidase crosslinking activity,” J. Biomed. Mater. Res. 66, 513–521 (2003). [CrossRef]
  11. T. S. Girton, T. R. Oegema, and R. T. Tranquillo, “Exploiting glycation to stiffen and strengthen tissue equivalents for tissue engineering,” J. Biomed. Mater. Res. 46, 87–92 (1999). [CrossRef] [PubMed]
  12. C. L. McIntosh, L. L. Michaelis, A. G. Morrow, S. B. Itscoitz, D. R. Redwood, and S. E. Epstein, “Atrioventricular valve replacement with the Hancock porcine xenograft: a five-year clinical experience,” Surgery 78, 768–775 (1975). [PubMed]
  13. H. Dardik, I. M. Ibrahim, R. Baier, S. Sprayregen, M. Levy, and I. I. Dardik, “Human umbilical cord. A new source for vascular prosthesis,” JAMA J. Am. Med. Assoc. 236, 2859–2862 (1976). [CrossRef]
  14. G. Wollensak, E. Spoerl, and T. Seiler, “Riboflavin/ultraviolet-A-induced collagen crosslinking for the treatment of keratoconus,” Am. J. Ophthalmol. 135, 620–627 (2003). [CrossRef] [PubMed]
  15. A. Jayakrishnan and S. R. Jameela, “Glutaraldehyde as a fixative in bioprostheses and drug delivery matrices,” Biomaterials 17, 471–484 (1996). [CrossRef] [PubMed]
  16. M. C. DeRosa and R. J. Crutchley, “Photosensitized singlet oxygen and its applications,” Coordin. Chem. Rev. 233–234, 351–371 (2002). [CrossRef]
  17. A. S. McCall, S. Kraft, H. F. Edelhauser, G. W. Kidder, R. R. Lundquist, H. E. Bradshaw, Z. Dedeic, M. J. C. Dionne, E. M. Clement, and G. W Conrad, “Mechanisms of corneal tissue cross-linking in response to treatment with topical riboflavin and long-wavelength ultraviolet radiation (UVA),” Invest. Ophthalmol. Visual Sci. 51, 129–138 (2010). [CrossRef]
  18. G. Wollensak, E. Spoerl, M. Wilsch, and T. Seiler, “Keratocyte apoptosis after corneal collagen cross-linking using riboflavin/UVA treatment,” Cornea 23, 43–49 (2004). [CrossRef] [PubMed]
  19. T. Tanabe, M. Oyamada, K. Fujita, P. Dai, H. Tanaka, and T. Takamatsu, “Multiphoton excitationevoked chromophore assisted laser inactivation using green fluorescent protein,” Nat. Methods 2, 503–505 (2005). [CrossRef] [PubMed]
  20. K. Koenig, I. Riemann, P. Fischer, and K. H. Halbhuber, “Intracellular nanosurgery with near infrared femtosecond laser pulses,” Cell Mol. Biol. (Paris) 45, 195–201 (1999).
  21. W. Denk, J. H. Strickler, and W. W. Webb, “Two-photon laser scanning fluorescence microscopy,” Science 248, 73–76 (1990). [CrossRef] [PubMed]
  22. D. Warther, S. Gug, A. Specht, F. Bolze, J. F. Nicoud, A. Mourot, and M. Goeldner, “Two-photon uncaging: new prospects in neuroscience and cellular biology,” Bioorgan. Med. Chem. 18, 7753–7758 (2010). [CrossRef]
  23. K. Kuetemeyer, R. Rezgui, H. Lubatschowski, and A. Heisterkamp, “Influence of laser parameters and staining on femtosecond laser-based intracellular nanosurgery,” Biomed. Opt. Express 1, 587–597 (2010). [CrossRef]
  24. P. K. Frederiksen, M. Jorgensen, and P. R. Ogilby, “Two-photon photosensitized production of singlet oxygen,” J. Am. Chem. Soc. 123, 1215–1221 (2001). [CrossRef] [PubMed]
  25. K. Koenig, “Multiphoton microscopy in life sciences,” J. Microsc. 200, 83–104 (2000). [CrossRef]
  26. A. Hopt and E. Neher, “Highly nonlinear photodamage in two-photon fluorescence microscopy,” Biophys. J. 80, 2029–2036 (2001). [CrossRef] [PubMed]
  27. S. Kalies, K. Kuetemeyer, and A. Heisterkamp, “Mechanisms of high-order photobleaching and its relationship to intracellular ablation,” Biomed. Opt. Express 2, 805–816 (2011). [CrossRef] [PubMed]
  28. A. Vogel, J. Noack, G. Huettman, and G. Paltauf, “Mechanisms of femtosecond laser nanosurgery of cells and tissues,” Appl. Phys. B 81, 1015–1047 (2005). [CrossRef]
  29. G. A. Blab, P. H. M. Lommerse, L. Cognet, G. S. Harms, and T. Schmidt, “Two-photon exciation action cross-sections of the autofluorescent proteins,” Chem. Phys. Lett. 350, 71–77 (2001). [CrossRef]
  30. W. R. Zipfel, R. M. Williams, R. Christie, A. Y. Nikitin, B. T. Hyman, and W. W. Webb, “Live tissue intrinsic emission microscopy using multiphoton-excited native fluorescence and second harmonic generation,” Proc. Natl. Acad. Sci. U.S.A. 100, 7075–7080 (2003). [CrossRef] [PubMed]
  31. R. A. Lorbeer, M. Heidrich, C. Lorbeer, D. F. Ramirez-Ojeda, G. Bicker, H. Meyer, and A. Heisterkamp, “Highly efficient 3D fluorescence microscopy with a scanning laser optical tomograph,” Opt. Express 19, 5419–5430 (2011). [CrossRef] [PubMed]
  32. M. H. Niemz, Laser-Tissue Interactions: Fundamentals and Applications (Springer, 2007).
  33. B. A. Roeder, K. Kokini, J. E. Sturgis, J. P. Robinson, and S. L. Voytik-Harbin, “Tensile mechanical properties of three-dimensional type I collagen extracellular matrices with varied microstructure,” J. Biomech. Eng. 124, 214–222 (2002). [CrossRef] [PubMed]
  34. G. Wollensak, E. Spoerl, and T. Seiler, “Stress-strain measurements of human and porcine corneas after riboflavinultraviolet-A-induced cross-linking,” J. Cataract Refractive Surg. 29, 1780–1785 (2003). [CrossRef]
  35. M. Eghbali and K. T. Weber, “Collagen and the myocardium: fibrillar structure, biosynthesis and degradation in relation to hypertrophy and its regression,” Mol. Cell. Biochem. 96, 1–14 (1990). [CrossRef] [PubMed]
  36. C. Xu and W. W. Webb “Measurement of two-photon excitation cross sections of molecular fluorophores with data from 690 to 1050 nm,” J. Opt. Soc. Am. B 13, 481–491 (1996). [CrossRef]
  37. B. P. Yu, “Cellular defenses against damage from reactive oxygen species,” Physiol. Rev. 74, 139–162 (1994). [PubMed]
  38. Z. H. Syedain, J. Bjork, L. Sando, and R. T. Tranquillo, “Controlled compaction with ruthenium-catalyzed photochemical cross-linking of fibrin-based engineered connective tissue,” Biomaterials 30, 6695–6701 (2009). [CrossRef] [PubMed]
  39. A. Vogel and V. Venugopalan, “Mechanisms of pulsed laser ablation of biological tissues,” Chem. Rev. 103, 577–644 (2003). [CrossRef] [PubMed]
  40. G. M. Fomovsky, J. R. Macadangdang, G. Ailawadi, and J. W. Holmes, “Model-based design of mechanical therapies for myocardial infarction,” J. Cardiovasc. Transl. Res. 4, 82–91 (2011). [CrossRef]

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