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Abstract
Doxorubicin (DOX) is a potent chemotherapeutic agent known to induce cardiotoxicity. Here we applied one-dimensional scanning multiphoton imaging to investigate the derangement of cardiac dynamics induced by DOX on a zebrafish model. DOX changed the cell morphology and significantly prolonged calcium transient and sarcomere contraction, leading to an arrhythmia-like contractile disorder. The restoration phase of calcium transient dominated the overall prolongation, indicating that DOX perturbed primarily the protein functions responsible for recycling cytosolic calcium ions. This novel finding supplements the existing mechanism of DOX cardiotoxicity. We anticipate that this approach should help mechanistic studies of drug-induced cardiotoxicity or heart diseases.
© 2021 Optical Society of America under the terms of the OSA Open Access Publishing Agreement
1. Introduction
Doxorubicin (DOX) is one of the most widely used chemotherapeutic drugs for cancer treatments [1,2]. It arrests the proliferation of cancer cells by binding to their DNA base pairs or topoisomerases [3–5]. Despite its anti-tumor efficacy, DOX induces multifaceted adverse effects to the heart (for instance, heart arrhythmia, decreased cardiac function, and life-threatening heart failure) in some populations of patients [1,2]. Cardiac complications of cancer chemotherapy not only reduce the quality of life but also increase the risk of death from cardiac-related causes, and therefore have limited the therapeutic potential of chemotherapy [6].
A previous study reported that DOX caused an unregulated release of calcium ions (Ca2+) from the cardiac sarcoplasmic reticulum (SR) [7]. Another study further showed that the elevated intracellular Ca2+ (or calcium overload) might cause degradation of proteins and lead to sarcomere disarray [8,9]. On the other hand, calcium homeostasis plays an essential role in regulating sarcomere contraction. Therefore, disturbance of calcium homeostasis would inevitably induce cardiac contractile dysfunction [10].
The zebrafish (Danio rerio) heart has a beating rate (about 120 beats per min) more comparable with human hearts than mice (between 500 to 800 beats per min). Furthermore, the cardiac action potential of zebrafish and humans shares main characteristics; besides, the inward and outward currents of their cardiac cells are qualitatively similar [11,12]. These features make zebrafish an attractive model for researches concerning cardiac electrophysiology.
Here we apply multiphoton imaging on a zebrafish model of chemotherapy-induced cardiotoxicity. This work aims to elucidate how DOX disturbs the calcium dynamics and subsequently interferes with cardiac contraction. To model non-acute cardiotoxicity, we injected DOX into adult zebrafish and imaged isolated cardiac cells three weeks after the injection. Using a rapid one-dimensional scanning (or “X-T” hereafter) mode, we simultaneously probed calcium transient and sarcomeric contraction on the same cardiac cells and revealed their temporal correlation.
We found that DOX deranged the morphology and sarcomeric structure of cardiac cells. Moreover, the cardiac calcium transient prolonged significantly, and these cardiac cells exhibited arrhythmia-like contractile behaviors. Importantly, our results reveal for the first time that the restoration phase of calcium transient dominated the overall prolongation, whereas the rising phase had insignificant change. Thus, we suggest that DOX perturbed mainly the protein functions responsible for the recycling of cytosolic Ca2+, which significantly extended the restoration phase of calcium transient. Subsequently, the prolonged calcium transient delayed sarcomere relaxation, thereby leading to arrhythmia-like contractile disorder. We envision that such an approach may be applicable for studying cardiotoxicity or heart diseases.
2. Materials and methods
2.1 Chemicals
Heparin, HEPES, taurine, glucose, calcium chloride, doxorubicin hydrochloride, and L-15 culture medium (Sigma Aldrich), collagenase II, collagenase IV and DPBS (Thermo Fisher Scientific), Primocin (Invivogen), Matrigel (Corning), and fetal bovine serum (HyClone) were purchased from the indicated sources.
2.2 Preparation of the zebrafish model of DOX-induced cardiotoxicity
The Animal Investigation Committee of the National Chiao Tung University (permit No. NCTU-IACUC-106005) approved the animal experiments.
Transgenic zebrafish, Tg(myl7:GCaMP), were purchased from Taiwan Zebrafish Core Facility at Academia Sinica (TZCAS) [13]. Adult fish aged between five to six months were used in this study. Figure 1(a) shows the timeline of protocols. Briefly, fish were fasted for twenty-four hours and then anesthetized with tricaine (0.75 mM). The experimental group was prepared by intraperitoneal injection of a DOX solution (20 mg/kg, 5 µL). This dose conforms to that used in a rodent model and is near the limiting dose for humans [14]. The control group was prepared in the same manner except that a DPBS solution of the same volume was used. After injection, fish were moved immediately to a tank with isolated running circulation and remained fasting for another twelve hours for recovery.
Fig. 1. Timeline and the multiphoton imaging setup of this study. (a) Timeline of the experiments. (b) Schematic of the home-built multiphoton microscope system. DM: dichroic mirror; GM: galvo mirrors; Obj: objective lens; CL: condenser lens; BF: bandpass filter; LF: low-pass filter; SHG: second-harmonic generation; TPEF: two-photon excited fluorescence.
2.3 Isolation and culture of primary ventricular cardiac cells
Cardiac cells were isolated from fish hearts three weeks after the injection (Fig. 1(a)) [15]. Before surgery, fish were anesthetized with tricaine (0.75 mM). Then their hearts were removed and placed in an iced heparin buffer (a DPBS solution containing 10 U/mL heparin and 100 µg/mL Primocin) to prevent the formation of blood clots. Next, ventricular tissues were carefully isolated from the hearts with a pair of sterile sharp forceps. Then, the isolated tissues were placed in a PBS solution (750 µL) containing HEPES (10 mM), taurine (30 mM), glucose (5.5 mM), CaCl2 (12.5 µM), and collagenases II and IV (5 mg/ml) for two hours maintaining at 29 °C in a thermomixer (ThermoMixer C, Eppendorf). After digestion, the solution was washed several times with a buffer (a DPBS solution containing 5% FBS, CaCl2 increased from 12.5 µM to 1000 µM) and centrifuged at 240 × g for 5 minutes. Next, cardiac cells were isolated and plated on a glass-bottomed dish precoated with Matrigel in an L-15 medium containing FBS (5%) and primocin (100 µg/mL). Finally, the dish was kept in an incubator maintaining at 28.5 °C until imaging experiments.
2.4 Multiphoton imaging of isolated cardiac cells
Imaging experiments were performed two days after cell isolation (Fig. 1(a)). The multiphoton microscope system was built by integrating a femtosecond-pulsed Ti:sapphire laser (Tsunami, Spectra-Physics) with a confocal scanner (FV300, Olympus) and an inverted microscope (IX71, Olympus) (Fig. 1(b)). To excite TPEF of GCaMP, the centeral wavelength of the laser was set at 900 nm [16]. The laser beam was focused on samples with a water immersion objective lens (60X, NA 1.2, Olympus). Two-photon excited fluorescence (TPEF) and second-harmonic generation (SHG) signals were collected in the epi direction with the same objective lens and in the forward direction with a condenser, respectively. Throughout imaging experiments, cardiac cells were maintained at 28.5 °C with a mini-incubator (MIU-IBC, Olympus) mounted on the microscope.
It has been shown that the intensity-squared profile of the illumination point-spread function provides a good estimation of the two-photon excitation volume and the 1/e widths of the lateral and axial intensity-squared profiles of the illumination are 175 nm and 451 nm, respectively, for an NA-1.2 objective lens at 900 nm [17]. Accordingly, the lateral and axial resolutions of our multiphoton imaging system are estimated to be 350 nm and 900 nm, respectively.
To stimulate cellular contraction, a square-pulse periodic electric field (Ep-p = 5 V/cm, 1 Hz, pulse-width = 30 ms) was applied to the medium with two sterile graphite electrodes. Dual-modal multiphoton (SHG and TPEF) imaging was first performed on cardiac cells. To determine calcium transient and sarcomere shortening, the confocal scanner was switched to a one-dimensional scanning (X-T) mode conducted along the contracting direction of cells (red lines, Fig. 2(a)). The scanning speed was approximately 500 lines per second (or 2.1 ms per scanning line) and the length of the one-dimensional scanning was 11.6 µm (512 pixels), respectively.
Fig. 2. Illustration of the protocol for determining sarcomeric shortening. (a) Representative TPEF and SHG images of isolated cardiac cells. After imaging, the scanner was switched to a rapid one-dimensional scanning (X-T) mode. The red line in the image highlights the one-dimensional scanning conducted along the contracting direction of sarcomeres. (b) Illustration of the sarcomere structure and the SHG profile obtained from the X-T mode imaging. (c) A representative SHG X-T image of a contracting cardiac cell triggered with a periodic stimulation (1 Hz). The red arrows indicate individual contracting events, whereas the white dots highlight the temporal position of individual sarcomere Z-lines. A cartoon of sarcomeres was displayed for illustration. (d) Trajectories of sarcomere Z-lines derived from the SHG X-T image. (e) Sarcomere shortening derived from the temporal change of sarcomere length. See texts for more details.
2.5 Image processing for the determination of the sarcomere shortening and the calcium transient
The sarcomere length is the distance between two adjacent sarcomere Z-lines, and the SHG signal is produced predominantly from the thick filament of sarcomeres [18]. Therefore, one can determine the sarcomere length from a distance between two adjacent minima along an SHG scanning line, as illustrated in Fig. 2(b). The temporal change of the sarcomere length was derived according to a protocol shown in Figs. 2(c-e). First, the intensity profile of individual SHG scanning lines was smoothed with local polynomial regression using MATLAB code. Next, the position of a sarcomere Z-line was identified from the local minima of each smoothed SHG scanning line (white dots in Fig. 2(c)). Then, the temporal trajectory of the Z-line was obtained by connecting the Z-line positions between adjacent scanning lines (red lines in Fig. 2(d)), and the trajectory was smoothed (dark lines in Fig. 2(d)). Next, the sarcomere length was determined from a distance between two adjacent Z-lines, and the sarcomere shortening was derived from the temporal change of the sarcomere length (Figs. 2(d-e)). Finally, the sarcomere contractility was determined by computing $({S{L_{relax}} - S{L_{contract}}} )/S{L_{relax}}$, where $S{L_{relax}}$ and $S{L_{contract}}$ denote the sarcomere length at the diastolic (relaxed) and systolic (contracted) states, respectively. Specifically, $S{L_{relax}}$ was determined by averaging the sarcomere length over 200 ms during the relaxation state.
To determine calcium transient, the TPEF signal produced from GCaMP was first summed along individual TPEF scanning lines. Then, calcium transient was determined by the temporal change of the added TPEF signal. Finally, the response of calcium transient was represented by $\Delta F/{F_0}$, where $\Delta F$ and ${F_0}$ denote the amplitude and basal value of calcium transient, respectively.
2.7 Statistics
Comparison of data from two groups was made with Student’s t-test (OriginPro, OiginLab). Values of p < 0.05 were considered statistically significant (* p < 0.05; ** p < 0.01; *** p < 0.001).
3. Results
3.1 Assessment of the cardioactivity of epinephrine
For demonstration, we first evaluated how a model cardioactive drug (epinephrine) modulated calcium transient and sarcomere contractility. Figure 3(a) demonstrates a representative result of TPEF/SHG X-T images acquired from isolated cardiac cells before and sixty seconds after treatment with epinephrine (100 nM). To assist in the comparison of the temporal relation between the calcium transient and the sarcomere contraction, we placed dashed lines with an interval of 1 sec at the onset of individual the rising TPEF signal.
Fig. 3. Demonstration of the pharmacological effect of epinephrine. (a) Representative TPEF/SHG X-T images acquired before and after an epinephrine treatment (100 nM, 60 s). The dashed lines were placed at the onset of the rising TPEF signal to assist in comparison of the temporal relation between the calcium transient and the sarcomere contraction. (b) Upper: calcium transient determined from the GCaMP signal. Lower: fractional change of calcium transient, defined as $\Delta F/{F_0}$ where $\Delta F$ and ${F_0}$ denote the amplitude and the basal value of calcium transient, respectively. (c) Upper: sarcomere shortening. Lower: sarcomeric contractility computed from $({S{L_{relax}} - S{L_{contract}}} )/S{L_{relax}}$ where $S{L_{relax}}$ and $S{L_{contract}}$ denote the sarcomere length at the relaxed (diastolic) and the contracted (systolic) states of cells, respectively.
The TPEF X-T images (green, Fig. 3(a)) show distinct contrasts synchronous with the change of the sarcomere length as highlighted by the dashed lines in the image. Comparison of the results obtained before and after the treatment further shows that the contrast of the image became more pronounced after the treatment. As shown in Fig. 3(b), the epinephrine treatment boosted the calcium transient of cardiac cells by a factor of 1.77 (* p < 0.05; n = 5).
In parallel, the SHG X-T images (red, Fig. 3(a)) show horizontal dark lines corresponding to temporal positions of individual sarcomere Z-lines. While this dose (100 nM, 60 s) did not change the contractile rhythm (dashed lines, lower panel, Fig. 3(a)), it increased the sarcomeric shortening (upper panel, Fig. 3(c)) and boosted the contractility by a factor of 1.61 (** p < 0.01; n = 5) (lower panel, Fig. 3(c)).
The above results are consistent with the pharmacological effect of epinephrine and demonstrate our approach’s ability to reveal a drug-induced change of calcium transient and sarcomere contractility of cardiac cells.
3.2 Assessment of the cardiotoxicity of DOX
Having demonstrated how a cardioactive drug modulated the calcium transient and sarcomere contractility of cardiac cells, we proceed to investigate non-acute cardiotoxicity induced by DOX with a protocol described in the experimental section.
Figure 4 demonstrates some representative TPEF (green) and SHG (red) images of cardiac cells isolated from adult zebrafish three weeks after the injection of DPBS (the control group) or DOX (the experimental group). The TPEF images (upper row, Fig. 4) reveal the morphology of cardiac cells. Specifically, the control group generally exhibited normal long rod-like morphology and had a smooth boundary. In contrast, the experimental group was much shorter and had a wavy boundary or even a protrusion. Furthermore, the SHG images (bottom row, Fig. 4) show that the control group exhibited orderly striated patterns of sarcomeres; in contrast, the experimental group showed discernible defects in sarcomeres. Together, these results show that DOX significantly deranged the morphology and sarcomere structures of cardiac cells.
Fig. 4. Representative images of cardiac cells (green: TPEF; red: SHG) obtained from the control and the experimental groups. Scale bar: 40 µm.
We next investigate how a DOX injection to adult zebrafish modulates the contractile dynamics of cardiac cells. Figure 5(a) demonstrates representative TPEF/SHG X-T images obtained from the control and the experimental groups. Specifically, cardiac cells of the control group exhibited periodic contractions, which were synchronous with the external stimulation at 1 Hz (upper, Fig. 5(a)). In contrast, cardiac cells of the experimental group had a significantly longer cycle relative to the control (lower, Fig. 5(a)). Specifically, these cardiac cells failed to keep up with the external stimulation's rhythm (1 Hz), and some contractions were missing. Notably, the contractions of these cardiac cells resembled an arrhythmia-like contractile disorder.
Fig. 5. Demonstration of DOX-induced cardiotoxicity. (a) Representative TPEF/SHG X-T images of cardiac cells isolated from the control (upper: DPBS treated) and the experimental groups (lower: DOX treated). The dashed lines were placed at the onset of the rising TPEF signal to assist in comparison of the temporal relation between the calcium transient and the sarcomere contraction. (b) Overlay of calcium transient (green) and sarcomere length (red) determined from the control group (upper) and the experimental group (lower). (c) Upper: overlay of calcium transient determined from the control (black line) and the experimental group (red line). Lower: Comparison of the duration of calcium transient, derived from the full width at half maximum of calcium transient. (d) Overlay of the time-varying sarcomere length determined from the control (black line) and the experimental group (red line).
Overlay of the calcium transient and sarcomere shortening further reveal their temporal correlation (Fig. 5(b)). In general, the rapid rising phase of calcium transient coincides with the drastic shortening of sarcomeres; so does the slower decreasing phase of calcium transient and the gradual relaxation of sarcomeres (upper, Fig. 5(b)). As described, the cycle of calcium transient and sarcomere contraction of the experimental group elongated significantly with a duration twice of the period of the external stimulation (lower, Fig. 5(b)).
To gain more mechanistic insight into the DOX-induced cardiac disorder, we overlaid the calcium transients of the control group and the DOX-treated group and displayed the result in Fig. 5(c). Quantitative analysis of calcium transient shows that the DOX treatment prolonged the duration of calcium transient (defined as the full width at half maximum of a calcium transient) by a factor of 3.85 (** p < 0.01; n = 6) (lower, Fig. 5(c)). Most strikingly, we found that the rising phase of both groups overlapped almost completely, whereas the decaying phase of the experimental group prolonged significantly relative to the control (upper, Fig. 5(c)). In other words, the result indicates that the DOX-induced prolongation of calcium transient occurred almost exclusively in the decaying phase.
Overlay of the sarcomere shortening between the control and the experimental groups shows a consistent feature. The temporal profiles of the shortening stage are comparable, whereas the relaxation stage of the experimental group exhibited elongated significantly compared to the control (Fig. 5(d)).
4. Discussion
Cardiac dysfunction or heart failure occurs in a subset of cancer patients treated with chemotherapy. Such cardiac complications not only limit the therapeutic potential of chemotherapy but also diminish patients’ quality of life [2]. For this reason, understanding and preventing drug-induced cardiotoxicity has attracted long-term research attention [19–21] and is a central focus of an emerging field termed cardio-oncology [6]. This study aims at elucidating how DOX, one of the most commonly used chemotherapeutic agents, interferes with cardiac dynamics. Using the rapid one-dimensional scanning mode of multiphoton imaging, we directly probed the temporal correlation between calcium transient and sarcomeric contraction. We show that DOX prolonged calcium transient and sarcomeric contraction, which subsequently perturbed cardiac cells’ rhythm and led to an arrhythmia-like contractile disorder. Importantly, we reveal for the first time that the decaying phase of calcium transient dominated the calcium transient prolongation (Fig. 5(c)), which caused a concurrent delay in the relaxation phase of sarcomeric contraction (Fig. 5(d)). We proposed a mechanism to explain the above findings.
Figure 6(a) illustrates the essential steps of cardiac excitation-contraction coupling. Briefly, an electrical stimulus triggers the voltage-gated opening of L-type calcium channels (LTCC) and then prompts a Ca2+ influx. The elevated cytosolic Ca2+ subsequently trigger “calcium-induced calcium release” from the SR through the ryanodine receptor (RyR), which later causes sarcomere contraction. Finally, the sarcoplasmic reticulum Ca2+-ATPase (SERCA) recycles the cytosolic Ca2+ back to SR, thus completing a cycle of excitation-contraction coupling. According to the mechanism, the rising and decaying phases of calcium transient are mainly regulated by RyR and SERCA, respectively. We have shown that a non-acute treatment of DOX in vivo significantly prolonged the calcium transient of cardiac cells. Remarkably, the decaying phase dominated the calcium transient prolongation (Fig. 5(c)). We note that the decaying phase of the calcium transient is associated with the recycling of cytosolic Ca2+ back to SR. Moreover, the calcium recycling is regulated mainly by SERCA, as illustrated in Fig. 6(a). Therefore, our result strongly indicates DOX (or its metabolites) impaired primarily the function of SERCA and subsequently led to the prolonged decaying phase of calcium transient.
Fig. 6. A proposed mechanism of DOX-induced cardiotoxicity. (a) Cartoon illustration of cardiac excitation-contraction coupling (LTCC: the L-type calcium channel; RyR: the ryanodine receptor; SERCA: the sarcoplasmic reticulum Ca2+-ATPase). (b) Cartoon illustration of a proposed mechanism of DOX-induced cardiotoxicity. See texts for more details.
While elucidating the mechanistic details requires more in-depth studies, numerous preceding works supported our deduction. First, a research group has reported that doxorubicinol (DOXol), a metabolite of DOX, can interact with SERCA [22]. Moreover, another two groups independently showed that DOX inhibited the transcription of SERCA [23,24]. Finally, another previous study found that treating cardiac cells with a SERCA inhibitor, thapsigargin, prolonged the restoration of cytosolic Ca2+ back to the basal level [25], a feature remarkably resembling our finding obtained on cardiac cells isolated from DOX-treated zebrafish. Figure 6(b) illustrates our proposed mechanism.
Although numerous previous studies have reported DOX-induced calcium overload and cell damage [26], we are the first, to the best of our knowledge, to show that DOX-induced disturbance occurred mainly in the restoration phase of calcium transient. We speculated that the difference in protocols might be responsible for why the subtle calcium derangement shown in Fig. 5 was not reported before. Specifically, we modeled non-acute cardiotoxicity by injecting DOX to adult zebrafish, and then we characterized calcium transient and sarcomere contraction three weeks post-injection. In comparison, it is common to induce cardiotoxicity by directly exposing cardiac cells to drugs in vitro. We suggest that these in vitro models might not recapitulate some pharmacological effects that occurred in vivo. For instance, it is generally not practical to treat cells for weeks; the limited exposure time might not be sufficient to induce non-acute toxic effects to cardiac cells. Besides, some preceding studies have reported that DOXol also contributed to DOX-induced cardiotoxicity [22]. More studies are necessary to confirm whether DOXol is produced in the in vitro model of DOX-induced cardiotoxicity.
Concerning biological models for cardio-electrophysiology or cardiotoxicity research, it is common to employ cardiac cells isolated from mice or the human-induced pluripotent stem cell-derived cardiomyocyte (hiPSC-CM). However, numerous preceding studies have shown that the cardiac electrophysiology, especially the action potential and electrocardiogram, between mice and human is notably different [27]. In contrast, the zebrafish is considered highly conserved in cardiac electrophysiology and cardiac Ca2+ signaling [11,28]. Compared to the hiPSC-CM model, a recent study has concluded that the zebrafish cardiovascular system provides a more reliable estimation of withdrawn cardiotoxic drugs, probably because of the immature electrophysiological phenotype of hiPSC-CMs [29,30].
A pioneering group of researchers has developed rapid one-dimensional scanning mode of multiphoton microscopy to study the alteration of microdomain Ca2+-contraction coupling on a rodent model of pressure-overload induced heart failure [31]. Specifically, they imaged calcium transient and sarcomeres by detecting TPEF from a fluorescent Ca2+ probe (fluo-4) and SHG from sarcomeres. Inspired by that study, we employed a similar approach to study DOX-induced cardiotoxicity. Particularly, we utilized transgenic zebrafish that express a genetic Ca2+ sensor (GCaMP) as a model animal [13,32]. The zebrafish has recently emerged as an alternative model for studying cardiac electrophysiology and cardiotoxicity because of its high degree of conservation in cardiac electrophysiology [11–14]. In addition, using transgenic zebrafish bearing GCaMP in the heart offers some convenience by eliminating the procedure of labeling cardiac cells with fluorescent probes.
5. Conclusion
In summary, we applied rapid one-dimensional scanning dual-modal multiphoton imaging on cardiac cells isolated from transgenic zebrafish hearts and revealed DOX-induced dynamic derangement of calcium transient and sarcomere contraction. Our novel finding supplements the existing mechanism of DOX cardiotoxicity. The combined use of one-dimensional scanning multiphoton imaging and transgenic zebrafish may become a valuable approach for studies concerning drug-induced cardiotoxicity and cardiac electrophysiology.
Funding
Center for Emergent Functional Matter Science of National Yang Ming Chiao Tung University (Ministry of Education, Taiwan); Ministry of Science and Technology, Taiwan (MOST 109-2113-M-009-026-MY2).
Acknowledgments
This work was supported by the Ministry of Science and Technology in Taiwan and the Center for Emergent Functional Matter Science of National Yang Ming Chiao Tung University from The Featured Areas Research Center Program within the framework of the Higher Education Sprout Project by the Ministry of Education (MOE) in Taiwan. The authors thank the support from Taiwan Zebrafish Core Facility (NSC101-2321-B400-014).
Disclosures
The authors declare no competing financial interests.
Data availability
Data underlying the results presented in this paper are not publicly available at this time but may be obtained from the authors upon reasonable request.
References
1. P. K. Singal and N. Iliskovic, “Doxorubicin-induced cardiomyopathy,” N. Engl. J. Med. 339(13), 900–905 (1998). [CrossRef]
2. C. Cristina, X. S. Renato, C. Susana, C. Sonia, J. O. Paulo, S. S. Maria, and I. M. Paula, “Doxorubicin: the good, the bad and the ugly effect,” Curr. Med. Chem. 16(25), 3267–3285 (2009). [CrossRef]
3. J. L. Nitiss, “Targeting DNA topoisomerase II in cancer chemotherapy,” Nat. Rev. Cancer 9(5), 338–350 (2009). [CrossRef]
4. O. Tacar, P. Sriamornsak, and C. R. Dass, “Doxorubicin: an update on anticancer molecular action, toxicity and novel drug delivery systems,” J. Pharm. Pharmacol. 65(2), 157–170 (2012). [CrossRef]
5. S. S. Tartakoff, J. M. Finan, E. J. Curtis, H. M. Anchukaitis, D. J. Couture, and S. Glazier, “Investigations into the DNA-binding mode of doxorubicinone,” Org. Biomol. Chem. 17(7), 1992–1998 (2019). [CrossRef]
6. C. G. Lenneman and D. B. Sawyer, “Cardio-oncology: an update on cardiotoxicity of cancer-related treatment,” Circ. Res. 118(6), 1008–1020 (2016). [CrossRef]
7. D. H. Kim, A. B. Landry, Y. S. Lee, and A. M. Katz, “Doxorubicin-induced calcium release from cardiac sarcoplasmic reticulum vesicles,” J. Mol. Cell. Cardiol. 21(5), 433–436 (1989). [CrossRef]
8. C. C. Lim, C. Zuppinger, X. Guo, G. M. Kuster, M. Helmes, H. M. Eppenberger, T. M. Suter, R. Liao, and D. B. Sawyer, “Anthracyclines induce calpain-dependent titin proteolysis and necrosis in cardiomyocytes,” J. Biol. Chem. 279(9), 8290–8299 (2004). [CrossRef]
9. B. Chen, L. Zhong, S. F. Roush, L. Pentassuglia, X. Peng, S. Samaras, J. M. Davidson, D. B. Sawyer, and C. C. Lim, “Disruption of a GATA4/Ankrd1 signaling axis in cardiomyocytes leads to sarcomere disarray: implications for anthracycline cardiomyopathy,” PLoS One 7(4), e35743 (2012). [CrossRef]
10. W. H. Barry and J. H. Bridge, “Intracellular calcium homeostasis in cardiac myocytes,” Circulation 87(6), 1806–1815 (1993). [CrossRef]
11. P. Nemtsas, E. Wettwer, T. Christ, G. Weidinger, and U. Ravens, “Adult zebrafish heart as a model for human heart? An electrophysiological study,” J. Mol. Cell. Cardiol. 48(1), 161–171 (2010). [CrossRef]
12. M. Vornanen and M. Hassinen, “Zebrafish heart as a model for human cardiac electrophysiology,” Channels 10(2), 101–110 (2016). [CrossRef]
13. N. C. Chi, R. M. Shaw, B. Jungblut, J. Huisken, T. Ferrer, R. Arnaout, I. Scott, D. Beis, T. Xiao, H. Baier, L. Y. Jan, M. Tristani-Firouzi, and D. Y. R. Stainier, “Genetic and physiologic dissection of the vertebrate cardiac conduction system,” PLoS Biol. 6(5), e109 (2008). [CrossRef]
14. X. Ma, Y. Ding, Y. Wang, and X. Xu, “A Doxorubicin-induced cardiomyopathy model in adult zebrafish,” JoVE 136(136), e57567 (2018). [CrossRef]
15. V. Sander, G. Suñe, C. Jopling, C. Morera, and J. C. I. Belmonte, “Isolation and in vitro culture of primary cardiomyocytes from adult zebrafish hearts,” Nat. Protoc. 8(4), 800–809 (2013). [CrossRef]
16. M. Drobizhev, N. S. Makarov, S. E. Tillo, T. E. Hughes, and A. Rebane, “Two-photon absorption properties of fluorescent proteins,” Nat. Methods 8(5), 393–399 (2011). [CrossRef]
17. W. R. Zipfel, R. M. Williams, and W. W. Webb, “Nonlinear magic: multiphoton microscopy in the biosciences,” Nat. Biotechnol. 21(11), 1369–1377 (2003). [CrossRef]
18. S. V. Plotnikov, A. C. Millard, P. J. Campagnola, and W. A. Mohler, “Characterization of the myosin-based source for second-harmonic generation from muscle sarcomeres,” Biophys. J. 90(2), 693–703 (2006). [CrossRef]
19. S. Zhang, X. Liu, T. Bawa-Khalfe, L.-S. Lu, Y. L. Lyu, L. F. Liu, and E. T. H. Yeh, “Identification of the molecular basis of doxorubicin-induced cardiotoxicity,” Nat. Med. 18(11), 1639–1642 (2012). [CrossRef]
20. A. Khalid, A. N. Mitropoulos, B. Marelli, S. Tomljenovic-Hanic, and F. G. Omenetto, “Doxorubicin loaded nanodiamond-silk spheres for fluorescence tracking and controlled drug release,” Biomed. Opt. Express 7(1), 132–147 (2016). [CrossRef]
21. J. Kress, D. J. Rohrbach, K. A. Carter, D. Luo, S. Shao, S. Lele, J. F. Lovell, and U. Sunar, “Quantitative imaging of light-triggered doxorubicin release,” Biomed. Opt. Express 6(9), 3546–3555 (2015). [CrossRef]
22. R. D. Olson, P. S. Mushlin, D. E. Brenner, S. Fleischer, B. J. Cusack, B. K. Chang, and R. J. Boucek, “Doxorubicin cardiotoxicity may be caused by its metabolite, doxorubicinol,” Proc. Natl. Acad. Sci. U.S.A. 85(10), 3585–3589 (1988). [CrossRef]
23. M. Arai, A. Yoguchi, T. Takizawa, T. Yokoyama, T. Kanda, M. Kurabayashi, and R. Nagai, “Mechanism of doxorubicin-induced inhibition of sarcoplasmic reticulum Ca2+-ATPase gene transcription,” Circ. Res. 86(1), 8–14 (2000). [CrossRef]
24. Y. H. Shih, Y. Zhang, Y. Ding, C. A. Ross, H. Li, T. M. Olson, and X. Xu, “Cardiac transcriptome and dilated cardiomyopathy genes in zebrafish,” Circ. Cardiovasc. Genet. 8(2), 261–269 (2015). [CrossRef]
25. A. Wrzosek, H. Schneider, S. Grueninger, and M. Chiesi, “Effect of thapsigargin on cardiac muscle cells,” Cell Calcium 13(5), 281–292 (1992). [CrossRef]
26. F. Timolati, T. Anliker, V. Groppalli, J.-C. Perriard, H. M. Eppenberger, T. M. Suter, and C. Zuppinger, “The role of cell death and myofibrillar damage in contractile dysfunction of long-term cultured adult cardiomyocytes exposed to doxorubicin,” Cytotechnology 61(1-2), 25–36 (2009). [CrossRef]
27. C. I. Berul, M. J. Aronovitz, P. J. Wang, and M. E. Mendelsohn, “In vivo cardiac electrophysiology studies in the mouse,” Circulation 94(10), 2641–2648 (1996). [CrossRef]
28. S. Kaese and S. Verheule, “Cardiac electrophysiology in mice: a matter of size,” Front. Physio. 3, 345 (2012). [CrossRef]
29. S. Dyballa, R. Miñana, M. Rubio-Brotons, C. Cornet, T. Pederzani, G. Escaramis, R. Garcia-Serna, J. Mestres, and J. Terriente, “Comparison of zebrafish larvae and hiPSC cardiomyocytes for predicting drug-induced cardiotoxicity in humans,” Toxicol. Sci. 171(2), 283–295 (2019). [CrossRef]
30. P. Machiraju and S. C. Greenway, “Current methods for the maturation of induced pluripotent stem cell-derived cardiomyocytes,” WJSC 11(1), 33–43 (2019). [CrossRef]
31. S. Awasthi, L. T. Izu, Z. Mao, Z. Jian, T. Landas, A. Lerner, R. Shimkunas, R. Woldeyesus, J. Bossuyt, B. M. Wood, Y.-J. Chen, D. L. Matthews, D. K. Lieu, N. Chiamvimonvat, K. S. Lam, Y. Chen-Izu, and J. W. Chan, “Multimodal SHG-2PF imaging of microdomain Ca2+-contraction coupling in live cardiac myocytes,” Circ. Res. 118(2), e19–e28 (2016). [CrossRef]
32. J. Nakai, M. Ohkura, and K. Imoto, “A high signal-to-noise Ca2+ probe composed of a single green fluorescent protein,” Nat. Biotechnol. 19(2), 137–141 (2001). [CrossRef]
