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Shock wave treatment (SWT) was shown to induce regeneration of ischaemic myocardium via Toll‐like receptor 3 (TLR3). The antimicrobial peptide LL37 gets released by mechanical stress and is known to form complexes with nucleic acids thus activating Toll‐like receptors. We suggested that SWT in the acute setting prevents from the development of heart failure via RNA/protein release. Myocardial infarction in mice was induced followed by subsequent SWT. Heart function was assessed 4 weeks later via transthoracic echocardiography and pressure–volume measurements. Human umbilical vein endothelial cells (HUVECs) were treated with SWT in the presence of RNase and proteinase and analysed for proliferation, tube formation and LL37 expression. RNA release and uptake after SWT was evaluated. We found significantly improved cardiac function after SWT. SWT resulted in significantly higher numbers of capillaries and arterioles and less left ventricular fibrosis. Supernatants of treated cells activated TLR3 reporter cells. Analysis of the supernatant revealed increased RNA levels. The effect could not be abolished by pre‐treatment of the supernatant with RNase, but only by a sequential digestion with proteinase and RNase hinting strongly towards the involvement of RNA/protein complexes. Indeed, LL37 expression as well as cellular RNA uptake were significantly increased after SWT. We show for the first time that SWT prevents from left ventricular remodelling and cardiac dysfunction via RNA/protein complex release and subsequent induction of angiogenesis. It might therefore develop a potent regenerative treatment alternative for ischaemic heart disease.
Myocardial ischaemia with consequent loss of cardiomyocytes can cause left ventricular remodelling including development of fibrotic scar tissue, thinning of the myocardium and decreased systolic function resulting in heart failure. Affected patients suffer from dyspnoea, fatigue and oedema, show high morbidity and mortality and represent a major socio‐economic health burden for Western countries 1, 2. Current treatment strategies provide symptomatic relief 3. However, no currently available treatment option is able to avoid ventricular remodelling with subsequent development of heart failure after myocardial injury.
Shock waves are mechanical pressure waves which have been used in medicine for kidney stone lithotripsy for more than 30 years 4. In lower energies, they were shown to induce tissue regeneration via induction of angiogenesis 5, mobilization of progenitor cells 6, 7 and alteration of inflammatory response 8. Therefore, they have been used successfully in clinical routine for regenerative purposes, for example in bone non‐unions 9, soft tissue wounds 10 and burn injuries 11. Cardiac SWT application showed induction of angiogenesis 5, improved function and caused symptom relief in patients suffering from chronic ischaemic heart disease 12. The exact working mechanism remains not fully understood.
We could describe only recently that the regenerative effects of SWT are mediated via RNA release and subsequent TLR3 activation. SW effects were missing completely in TLR3−/− animals 13. However, extracellular self‐RNA is degraded immediately to avoid immunogenic response 14, 15. Therefore, it remains unknown how extracellular RNA released upon SWT (1) escapes from degradation and (2) can activate intracellular TLR3.
Extracellular RNA mainly occurs in protein‐complexed forms, and thus escapes degradation 16. Thereby, the cationic antimicrobial peptide LL37 has been described to be released upon mechanical stress, to form complexes with ribonucleic acids, to pass the cellular membrane and to subsequently activate intracellular TLRs 15. We therefore suggested that SWT causes release of LL37. The peptide forms complexes with extracellular RNA, passes the cell membrane and activates TLR3.
In this study, we aimed to investigate whether (1) SWT in the acute setting of myocardial infarction prevents from the development of ischaemic heart failure, and (2) SWT effects are indeed mediated via LL37/RNA complexes.
The experiments were approved by the institutional animal care and use committee at Innsbruck Medical University and by the Austrian ministry of science. The investigation conformed to the ‘Guide for the Care and Use of Laboratory Animals’ published by the U.S. National Institutes of Health (NIH Publication No. 85‐23, revised 1996; available from: www.nap.edu/catalog/5140.html). Male, 12‐ to 14‐week old C57/BL6 mice (Charles River, Sulzfeld, Germany) weighing 25–30 g were randomly divided into two groups (shock wave therapy group = SWT and untreated control group = CTR, n = 6). SWT animals received shock wave therapy directly after LAD ligation. CTR animals were left untreated. The animals were killed 4 weeks after therapy for harvesting of the heart.
Myocardial infarction was induced as it is the standard model for the induction of ischaemic cardiomyopathy. It was performed as described previously 17. Briefly, animals were anesthetized by an intraperitoneal injection of ketamine hydrochloride (Graeub, Bern, Switzerland; 80 mg/kg bodyweight) and xylazine hydrochloride (aniMedica, Senden, Germany; 5 mg/kg bodyweight). A left lateral thoracotomy was performed in the fourth intercostal space for exposure of the heart. The left anterior descending artery (LAD) was ligated at the level of the pulmonary artery using 7‐0 polypropylene sutures (Ethicon, Somerville, NJ, USA).
Animals were anesthetized for SWT as described above. Commercially available ultrasound gel was used for coupling. The commercially available Orthogold 180 with the hand‐held applicator CG050‐P (TRT LLC, Tissue Regeneration Technologies, Woodstock, GA, USA), which was specifically designed for cardiac shock wave therapy, was used for the treatment. Three‐hundred impulses were delivered to the heart through the thorax aiming at the ischaemic area at an energy flux density of 0.38 mJ/mm2 at a frequency of 5 Hz. At these energy levels, no adverse effects could be observed. However, for all in vitro experiments, 250 impulses with an energy flux density of 0.08 mJ/mm2 at a frequency of 3 Hz were used. 17 The rationale of the treatment parameters is our experience from previous studies 18.
Haemodynamic pressure–volume analysis and transthoracic echocardiography were performed, as the combination of these methods allows exact measurement of cardiac function. Invasive haemodynamic measurements and the analysis of pressure–volume (PV) loops were performed as terminal procedure according to established protocols described previously 18 using a pressure‐volume conductance system (MPVS Ultra; Millar Instruments, Houston, TX, USA) connected to the PowerLab 8/35 data acquisition system, United Kingdom, and analysed using LabChart7pro software system (AdInstruments, Colorado Srpings, CO, USA), United Kingdom. Mice were anaesthetized (3% isoflurane and 96–97% O2), intubated and ventilated by a small rodent ventilator. The animals were placed on a temperature‐controlled heating platform, and the core temperature was maintained at 37.5°C. Anaesthesia was reduced and kept at 1.5% isoflurane and 98.5% O2. A polyethylene catheter was inserted into the right external jugular vein for saline calibration (10% NaCl) at the end of the experiment to calculate the parallel volume. The 1.4‐F PV conductance catheter (SPR‐839; Millar) was inserted into the right carotid artery and advanced into the ascending aorta. Aortic (systemic) blood pressure and heart rate were recorded after stabilization for 5 min. Then, the catheter was advanced into the LV under pressure control through the aortic valve. Parameters of systolic and diastolic function, including LV end‐diastolic pressure (LVEDP), LV end‐systolic volume (LVESV), LV end‐diastolic volume (LVEDV), ejection fraction (EF), maximal slope of LV systolic pressure increment (dP/dt max) and maximal slope of diastolic pressure decrement (dP/dt min), were measured and calculated according to standard formulas. After baseline measurements, transient occlusion of the inferior vena cava was performed and used to derive end‐systolic (Ees) and end‐diastolic pressure–volume relationships (EDPVR) as load‐independent measures of cardiac contractility and relaxation, respectively. Heart rate (HR) was separately measured via surface ECG electrodes.
Transthoracic echocardiography was performed using standardized protocols for the assessment of heart function and morphometry as described previously 19, 20. Briefly, lightly anaesthetized mice (0.5% isoflurane and 99.5% O2) were placed on a temperature‐controlled warming pad (kept at 37.5°C) and imaged in the supine position using a high‐resolution micro‐imaging system equipped with a 30‐MHz linear array transducer (Vevo770TM Imaging System; VisualSonics Inc., Toronto, Ontario, Canada), respectively. Standard 2D‐ and M‐mode tracings of the left ventricle (LV; long axis and short axis at papillary muscle level) were recorded, and ejection fraction was averaged from three consecutive cardiac cycles under stable conditions. M‐Mode pictures were recorded in the mid‐papillary portion as standard in the Teichholz measuring technique. The investigator was blinded to the treatment.
Cell culture and in vitro assays were performed to investigate underlying mechanisms of SW‐induced angiogenesis. For this purpose, human umbilical vein endothelial cells (HUVECs) were cultured as described previously 21. Permission was given from the ethics committee of Innsbruck Medical University (No. UN4435). Cells were used until passage 5. All in vitro experiments were performed in triplicates.
In all experiments, 250 impulses were applied to the SWT group at an energy flux density of 0.08 mJ/mm2 and a frequency of 3 Hz. These parameters were chosen due to our experience from previous experiments 21, 22.
For assessment of proliferation, a BrdU assay (Roche, Rotkreuz, Switzerland) was performed as recommended by the manufacturer. Tube formation assay was performed as described previously 23. Tube formation was analysed using the ImageJ plugin Angiogenesis Analyzer as described previously 17. Both assays are recognized assays for the assessment of in vitro angiogenesis. For reporter cell assays, a HEK reporter cell line (TLR3/ISRE LUCPorter; Imgenex, San Diego, CA, USA) was purchased and used as suggested by the manufacturer. Cells were treated with SWT as described above. 24 hrs after treatment, cells were washed with PBS and luminescence was analysed using a Luciferase Assay System (Promega, Fitchburg, WI, USA).
RNA content was measured as described previously, as it represents a feasible technique for RNA content measurement 13. Rhodamine labelled polyinosinic:polycytidylic acid (poly (I:C)) (Invivogen, San Diego, CA, USA) was added to HUVEC supernatant prior to SWT treatment. Twenty‐four hours after, SWT cells were washed with PBS, cell membranes were stained using wheat germ agglutinin (WGA; Life Technologies, Carlsbad, CA, USA) and nuclei were visualized using DAPI. Rhodamine fluorescence was quantified using ImageJ. This method has proven useful for the investigation of RNA uptake 15. Results are depicted as positive area.
Western blot for protein expression was performed as described previously 13. The blots were probed with LL‐37 antibody (Abcam, Cambridge, UK).
Immunofluorescence staining was performed as described previously to analyse proliferation and number of vessels 13. Histological sections of the heart were incubated with monoclonal rat anti‐CD31 (nova, Hamburg, Germany) rabbit polyclonal anti‐alpha smooth muscle actin antibodies (Abcam), rabbit anti‐Ki 67 or rabbit anti‐LL37 over night at 4°C. Masson–Goldner trichrome staining was performed as suggested by the manufacturer (Carl Roth GmbH, Karlsruhe, Germany) to quantify post‐infarctional fibrosis. The area of fibrosis and total area of the left free ventricle were measured using ImageJ (NIH, Bethesda, MD, USA). Values are shown as the ratio of fibrotic area to total left ventricular free wall as described previously 24. Five areas per sample were analysed. Three random pictures per sample were analysed. Sections were examined with a Zeiss Axioplan 2 (Zeiss, Oberkochen, Germany) and a Leica SP5 confocal microscope (Leica, Wetzlar, Germany). Images were analysed using ImageJ software (National Institutes of Health) and processed with Adobe Photoshop CS5.1 for Mac (Adobe Systems Inc., San Jose, CA, USA). Analyses were performed by a single blinded researcher.
A LL‐37 ELISA kit was used to analyse LL37 release to the supernatant (Hycult Biotech, Uden, the Netherlands). The assay was performed as recommended by the manufacturer.
All results are expressed as mean + S.E.M. Statistical comparisons between two groups were performed by Student's t‐test or Mann–Whitney U‐test as appropriate. Multiple groups were analysed by two‐way anova followed by Bonferroni's multiple comparison test to determine statistical significance. Probability values <0.05 were considered statistically significant. All experiments were repeated at least in triplicate.
The design of the in vivo experiment is depicted in Figure Figure1(A).1(A). Baseline echocardiography showed no difference between the groups before treatment (ejection fraction (EF) in %: CTR 45.25 ± 1.11 versus SWT 46.2 ± 0.66, P = 0.47; Fig. Fig.1B).1B). Four weeks after induction of myocardial infarction and subsequent shockwave treatment, transthoracic echocardiography showed significantly improved ejection fraction in treated animals compared with untreated control animals (LVEF in %: CTR 35.25 ± 1.11 versus SWT 46 ± 2.83, P = 0.0122; Fig. Fig.1B1B and C). This finding could be confirmed by haemodynamic pressure‐volume measurement (LVEF in %: CTR 36.25 ± 1.70 versus SWT 45.25 ± 2.43, P = 0.0029; Fig. Fig.1D1D and E). In addition, a decreased left ventricular end‐diastolic pressure (LVEDP) following SWT was found (Fig. (Fig.1F;1F; mmHg: CTR 15.75 ± 0.85 versus SWT11.75 ± 1.38, P = 0.0485). Myocardial contractility was increased in treated animals (Fig. (Fig.1G1G and I; dP/dt max in mmHg/sec.: CTR 6222 ± 157.7 versus SWT 7236 ± 313.5, P = 0.0276; dP/dt min in mmHg/sec.: CTR −5906 ± 368 versus SWT −6396 ± 289, P = 0.3354). Moreover, left ventricular end‐diastolic volume (Fig. (Fig.1H;1H; LVEDV) was decreased and left ventricular end‐systolic volume (LVESV) was increased after SWT, however, not significantly (Fig. (Fig.1J;1J; LVEDV in μl: CTR 22.25 ± 2.78 versus SWT 19.256 ± 1.49, P = 0.3785; LVESV in μl: CTR 14.25 ± 2.10 versus SWT 10.5 ± 0.65, P = 0.1382).
In a next step, we aimed to clarify whether SWT resulted in angiogenesis and reduction in the fibrotic scar (Fig. (Fig.2A).2A). Treated hearts showed significantly increased numbers of capillaries (Fig. (Fig.2B;2B; capillaries per HPF: CTR 1.5 ± 0.23 versus SWT 7.17 ± 0.76, P < 0.0001) and arterioles (Fig. (Fig.2C;2C; arterioles per HPF: CTR: 1.1 ± 0.21 versus SWT 4.23 ± 0.32, P < 0.0001). In addition, we found significantly decreased amounts of fibrosis after SWT (% of free LV: CTR ± 2.76 versus SWT 8.97 ± 3.08, P = 0.0132; Fig. Fig.2D2D and E).
Endothelial cells were treated with SWT and analysed for proliferation (Fig. (Fig.3A).3A). We found increased proliferation after treatment (Fig. (Fig.3B;3B; % of Ki67‐positive cells: CTR 2.46 ± 1.25 versus SWT 12.51 ± 3.10, P = 0.0047). Next, we aimed to clarify whether the observed effect was due to a factor that was released into the supernatant. Treatment with pre‐conditioned medium indeed resulted in higher proliferation rates (compared to cells treated with unconditioned medium; Fig. Fig.3C;3C; arbitrary units: CTR 0.07 ± 0.01 versus SWT 1 hr 0.07 ± 0.003, P = 0.28; SWT 6 hrs 0.11 ± 0.01, P = 0.0019 versus CTR; SWT 24 hrs 0.07 ± 0.004, P = 0.86 versus CTR). Next, we analysed the RNA content in treated supernatants. We found a significantly increased amount of total RNA in supernatants after SWT compared to untreated controls (ng/ml: CTR 20.21 ± 2.56 versus SWT 47.7 ± 6.97, P = 0.0023; Fig. Fig.3D3D and E). Addition of pre‐marked RNA to cell culture supernatants prior to SWT showed an increased uptake 3 hrs after treatment (Fig. (Fig.3F3F and G; arbitrary units: 1 hr: CTR 1855 ± 824.7 versus SWT 5007 ± 1061, P = 0.0625; 3 hrs: CTR 31.67 ± 28.17 versus SWT 19,757 ± 1054, P = 0.0001; 6 hrs: CTR 545.7 ± 81.62 versus SWT 453.3 ± 44.67, P = 0.3722; 24 hrs: CTR 10,387 ± 1261 versus SWT 8249 ± 3862, P = 0.6267).
Next, we added either RNase or proteinase and a combination of both to the endothelial cells prior to SWT and again analysed for proliferation. RNase alone did not abolish the SWT effect (arbitrary units: CTR 0.22 ± 0.08, SWT 0.48 ± 0.04, P = 0.02 versus CTR; SWT + RNase 0.52 ± 0.06, P = 0.02 versus CTR). However, SW effects were abolished in the presence of proteinase (SWT + Proteinase 0.17 ± 0.03, P = 0.5312 versus CTR; Fig. Fig.4A).4A). The results were confirmed in a tube formation assay showing increased number of segments (Fig. (Fig.4B;4B; segments per HPF: CTR 320.8 ± 13.78 versus SWT 462.5 ± 22.26, P = 0.0015), junctions (Fig. (Fig.4C;4C; junctions per HPF: CTR 263.5 ± 9.82 versus SWT 369.7 ± 16.4, P = 0.0013) and nodes (Fig. (Fig.4D;4D; nodes per HPF: CTR 891.3 ± 25.96 versus SWT 1292 ± 60.11, P = 0.0029) after SWT. The effects could not be reversed by addition of RNase, only treatment with additional proteinase resulted in abolished SW effects (segments per HPF: SWT+RNase: 440 ± 31.42, P = 0.0186 versus CTR; SWT+Proteinase: 208 ± 73.37, P = 0.136 versus CTR; SWT+Proteinase+RNase: 170 ± 16.56, P = 0.0009 versus CTR; junctions per HPF: SWT+RNase: 354.2 ± 21.29, P = 0.0113; SWT+Proteinase: 182 ± 53.03, P = 0.1357 versus CTR; SWT+Prot+RNase: 159 ± 12.11, P = 0.0005 versus CTR; nodes per HPF: SWT+RNase 1246 ± 77.42, P = 0.0175 versus CTR; SWT+Proteinase 572 ± 170.9, P = 0.1771 versus CTR; SWT+Proteinase+RNase 554.8 ± 29.93, P = 0.0005 versus CTR).
LL37 expression was increased after SWT as shown by Western blot analysis (Fig. (Fig.5A5A and B; arbitrary units: CTR 40.35, SWT 30’ 39.13, SWT 1 hr 29.15, SWT 2 hrs 25.25, SWT 5 hrs 49.14, SWT 24 hrs 61.18, SWT 48 hrs 77.59) as well as immunofluorescence analysis (Fig. (Fig.5C5C and D; arbitrary units: CTR 47,496 ± 10,690 versus SWT 139,243 ± 22,250, P = 0.0054). In addition, LL37 content in the supernatant was also increased after SWT (Fig. (Fig.5E;5E; ng/ml: CTR 1.17 ± 0.0005 versus SWT 0 hr 1.65 ± 0.008, P < 0.0001, SWT 2 min. 1.30 ± 0.006, P < 0.0001 versus CTR; SWT 1 hr 1.41 ± 0.002, P < 0.0001 versus CTR; SWT 6 hrs 1.52 ± 0.008, P < 0.0001 versus CTR; SWT 24 hrs 1.32 ± 0.005, P < 0.0001).
Finally, we transferred pre‐treated supernatant onto TLR3 reporter cells. Supernatant from SW‐treated cells caused TLR3 activation (arbitrary units: CTR 8728 ± 3138 versus SWT 131,874 ± 12,328, P < 0.0001). This effect could again not be abolished by pre‐treatment with RNase (arbitrary units: SWT+RNase 110,171 ± 14,482, P = 0.0005 versus CTR), but only with additional proteinase treatment (arbitrary units: SWT+Prot+RNase 877 ± 380.8, P = 0.089 versus CTR) or sole proteinase treatment (arbitrary units: SWT+Prot 18,713 ± 11,018, P = 0.46 versus CTR; Fig. Fig.5F)5F) clearly indicating the pivotal role of a RNA/protein complex in SW‐induced TLR3 stimulation.
Heart failure due to ischaemic heart disease represents a global socio‐economic burden 25. Currently, there are no treatment options available for preventing its development after myocardial infarction. SWT represents a promising therapeutic tool for the treatment and regeneration of ischaemic myocardium 5. In the present study, we treated hearts after induction of myocardial infarction and evaluated cardiac function 4 weeks later. Untreated animals developed poor cardiac function with low ejection fraction, impaired contractility and deteriorated systolic and diastolic function. These results could be shown in transthoracic echocardiography and confirmed in haemodynamic pressure–volume measurements. However, functional analyses showed that left ventricular function and myocardial contractility were significantly improved after SWT. In addition, treated animals showed significantly decreased amount of fibrosis of the left ventricle after SWT. These results show clearly a protective effect of SWT concerning ventricular remodelling and cardiac function after myocardial infarction.
Cardiac regeneration after chronic ischaemia has been shown in small and large animal models via induction of angiogenesis 5, 12, 26. In addition, cardiac SWT caused improvement of symptoms in patients suffering from chronic ischaemic heart disease. However, we show for the first time that SWT after acute myocardial infarction attenuates the decrease in cardiac function concerning contractility, diastolic and systolic function.
We found increased numbers of capillaries as well as arterioles in treated myocardium. This might be a hint that the protective effect of SWT after myocardial infarction is due to early induction of angiogenesis. The fact that we found increased numbers of arterioles shows that SWT indeed induces the formation of long‐term stable vessels.
Our group recently showed that the SW effect is mediated via TLR3 13. However, it remains unclear how the receptor is activated upon mechanical stimulation with SWT. Supernatant from cells pre‐conditioned with SWT showed similar effects on proliferation as the direct application of SWT to cells does. This gave us a strong hint that whatever causes the SWT effect is released into the supernatant. TLR3 is known to be activated by RNA 27. We therefore measured RNA content in the supernatants and could show that it is indeed increased after SWT. However, it remained unclear how the extracellular RNA escapes degradation and passes the cellular membrane to activate intracellular TLR3. The fate of RNA is known to be determined by RNA‐binding proteins 16. We therefore suggested that released RNA binds to a protein and thus passes the cellular membrane. To test this hypothesis, we applied SWT to a tube formation assay in the presence of RNase and proteinase. Interestingly, RNase treatment did not abolish the SWT effect, but only additional treatment with proteinase did indicating the existence of a RNA/protein complex.
In a next step, we aimed to specify which protein is responsible for the observed effects. Gilliet et al. have identified LL37 as a key player in the development of psoriasis. Thereby, this antimicrobial peptide gets released upon mechanical stimulation, binds nucleic acids and activates intracellular nucleic acid TLRs 28. We analysed treated cells for LL37 release and LL37 expression and found both increased after SWT. When we added the pre‐marked RNA analogon Poly(I:C) to cells prior to treatment, we found significantly increased uptake of RNA after SWT. These results hint towards involvement of LL37 after SWT. However, it remains unknown whether other proteins/RNA complexes are also involved.
Finally, we analysed whether TLR3 activation after SWT was indeed accomplished by protein/RNA complexes. For this purpose, we performed a TLR3 reporter cell assay. SWT leads to the activation of TLR3, which could not be abolished by RNase treatment, but only with additional proteinase treatment showing that indeed RNA/protein complexes lead to TLR3 activation after SWT.
Summarizing, we could show for the first time that SWT after acute myocardial infarction prevents from left ventricular remodelling and cardiac dysfunction via induction of angiogenesis. In addition, we found that the cellular response upon SWT is mediated via RNA/protein complexes with involvement of the antimicrobial peptide LL37 activating TLR3. SWT could develop a potent therapeutic tool for the treatment of acute myocardial ischaemia and the prevention of ischaemic heart failure.
None. The authors confirm that there are no conflicts of interest. The sponsors of this study had no role in study design, data collection, analysis and decision to publish or prepare the manuscript.
C.T. and J.H. designed the experiments and wrote the manuscript. C.T., M.G., U.P. and D.L. performed animal experiments. C.T., L.P. and E.K. performed in vitro experiments. C.T., U.P., E.K., E.J.P. and F.N. performed analyses. M.G. revised the manuscript.
We kindly thank Annabella Knab for performing tissue sections. This work was in part supported by a research grant of Medizinischer Forschungsfonds Tirol (MFF) [project no. 220] and by a research grant provided by TRT—Tissue Regeneration Technologies LLC., Woodstock, Georgia, USA, both to J.H.