With thanks to Natalie Johnston for the recording of the non-surgical footage and all video editing.

All procedures described in this experiment were approved by the Animal Ethics Committee at the Northern Sydney Local Health District, NSW, Australia (project number RESP17/55).
1. Anesthesia and intubation
NOTE: Turn on and set up the stereomicroscope, the heat pad (covered with an absorbant sheet) and ventilator system.
<ol>
	<li>Clean gloves, the surgical area, and the tools with 70% ethanol.</li>
	<li>Weigh the mouse to calculate the dosage of anesthesia injected by the intraperitoneal route (ketamine 40 mg/kg, xylazine&#160;5&#8201;mg/kg, atropine&#160;0.15&#8201;mg/kg) and give the injection.</li>
	<li>Once the mouse reaches a deep plane of anesthesia, shave the ventral left side of the thorax with a trimmer.</li>
	<li>Place the mouse in a chamber containing 2% isoflurane (ensuring adequate extraction ventilation in the room). 
	NOTE: The relatively low dose of ketamine/xylazine injection together with 2% isoflurane inhalation reduces the risk of mouse death while allowing optimal intubation without waking the mouse up.</li>
	<li>Place the mouse supine and restrain it from its upper incisor teeth with a 3.0 suture taped to the bench, as shown in the video. Confirm sedation by performing a toe pinch. Position a high-intensity illuminator above the mouse neck so that the oropharynx can be visualized. 
	NOTE: Alternatively, the mouse can be placed on the stand from the intubation kit (e.g., Kent Mouse Intubation Kit) with an elastic band secured under the top incisors to hold the mouth open to identify the trachea.</li>
	<li>Use a curved spatula to open the jaw and another pair of spatulas/blunt forceps to lift the tongue gently out of the way. Be sure to intubate while positioned at or slightly below eye-level with the body of the mouse.</li>
	<li>Visualize the opening and closing of the vocal cords. When open, insert the 20 G plastic catheter supplied with the intubation kit.</li>
	<li>Carefully transfer the intubated mouse to an operating surface equipped with a heating pad. Connect the mouse to the ventilator (e.g., MouseVent) that automatically sets the target volume based on mouse weight.</li>
	<li>Deliver 1.5-2% isoflurane with oxygen (which is automatically regulated by the ventilator: ensure there is a connection from an oxygen cylinder to the automatic ventilator at 1-2 L/min flow rate to the ventilator). Verify the intubation by checking for bilateral chest rise. Verify anesthesia by performing a toe pinch.</li>
	<li>Apply opthalmic ointment (e.g., Puralube Vet Opthalmic Ointment) to both eyes to prevent them from drying out.</li>
</ol>
2. Preparing the surgical field
<ol>
	<li>Secure the intubation tube with tape at the connecting site between the ventilator and the breathing tube/catheter.</li>
	<li>Cut a longer piece of tape and secure its left front foot to the operating surface in a slightly elevated position. Also tape down the other extremities.</li>
	<li>Clean the chest with sterile 70% isopropanol and povidone iodine solution, cleaning in a circular motion moving from center to periphery.</li>
	<li>Verify anesthesia once more with a toe pinch.</li>
	<li>Administer 0.08 mg/kg Temvet (buprenorphine) in 0.1 mL of 0.9% saline via subcutaneous injection.</li>
</ol>
3. Left lateral thoracotomy
<ol>
	<li>Use fine-tip forceps to gently lift the skin at a point approximately 5 mm to the left of the prominent xiphoid cartilage. Use surgical scissors to create a superomedial incision in the skin from this point upwards and towards the midline, to the level of the manubrium.</li>
	<li>Use curved forceps to gently separate the skin and muscle layers. Open the muscle layer, following the skin incision.</li>
	<li>Identify and make an incision in the third intercostal space, following the natural angle of the ribcage.</li>
	<li>Use a retractor to gently spread apart the 3rd and 4th ribs.</li>
	<li>Gently remove the thin pericardium with forceps.</li>
	<li>If the LAD is not visualized, gently push the left auricle (see Supplementary Figure 1) upwards and locate the coronary arteries underneath.</li>
</ol>
4. Left anterior descending (LAD) permanent coronary artery ligation
<ol>
	<li>Cut a ~3 mm long 3-0 silk suture and put this reinforcing 3-0 silk suture piece on top of the LAD in the same direction as the LAD (as shown in the video at time point 02:12 &#8211; 02:20).</li>
	<li>Identify the LAD and pass a 7-0 silk suture under the LAD. If the LAD is not clearly visualized, insert the needle 1 mm inferior and medial to the inferiormost point reached by the tip of the left auricle during dynamic movement of the heart. 
	NOTE: This structure is a lighter color red to the ventricular chambers of the heart but darker than the adjacent lung and is best visualized in the video at time point 01:54 &#8211; 01:55 where it is visible just inferior to the superior arm of the retractor, superior to the left lung (see Supplementary Figure 1 for annotated video still image).</li>
	<li>Complete two throws with the 7-0 silk suture and close it tightly passing on top of the supportive 3-0 silk suture to secure the LAD. If the ligation is successful, the anterior ventricular area distal from the ligature will blanch.</li>
	<li>Complete the knot with a third throw in the opposite direction to secure it, ensuring no upward traction force is transmitted to the suture. Additional throws are not required to reduce the risk of damage to the myocardium or LAD by suture cutting through.</li>
</ol>
5. Transplantation of the bioprinted patch onto the epicardium
<ol>
	<li>Carefully move the bioprinted patch from a six well plate to the infarct area by using the sterile inside surface of an opened surgical scalpel packet.</li>
	<li>Carefully position the bioprinted patch onto the anterior epicardial surface, where it should cover the whole surface and drape over the inferior and lateral edges, covering the left ventricle and the infarct zone (blanched area).</li>
	<li>Gently close and remove the retractor without directing sharp edges towards the heart.</li>
	<li>Use 6-0 prolene sutures in a simple interrupted pattern to close the ribcage and the muscle layers.</li>
	<li>With the Sigh Breath function while closing the chest with the 6-0 prolene sutures, inflate the lungs to remove excess air in the pleural cavity, which would otherwise become trapped in the chest cavity and result in a pneumothorax.</li>
	<li>Ensure that the chest is tightly sealed.</li>
	<li>Decrease the isoflurane to 1.0%. Close the skin with 6-0 prolene sutures in a simple interrupted pattern. Turn the isoflurane vaporizer off.</li>
</ol>
6. Mouse recovery
<ol>
	<li>Topically apply 2 mg/mL bupivacaine in 0.9% saline to the incision. Administer also: i) Antisedan (atipamezole) 1 mg/kg; ii) Lasix (furosemide) 8 mg/kg; iii) 600 &#181;L of 0.9% saline solution via a subcutaneous injection. 
	NOTE: Antisedan is to reverse the anesthetic more rapidly; furosemide is to offload excess fluid due to cardiac output compromise and additional fluid administered with drug injections.</li>
	<li>Monitor the mouse and wait until independent breathing is observed to remove the mouse from the intubation tube.</li>
	<li>When the mouse demonstrates an adequate bilateral breathing rate and depth and responds to a toe pinch, place the mouse in a clean recovery cage placed on a heat pad.</li>
	<li>Provide the mouse with moist food (moistened for chewability), a water bottle and nutrient/hydrating gel. Monitor for an exaggerated breathing effort, excessive bleeding, or other potentially life-threatening complications.</li>
	<li>For the next three days, administer 0.08 mg/kg Temvet (buprenorphine) in 0.1 mL of 0.9% saline via subcutaneous or intraperitoneal injection, twice daily, then once daily up to the fifth day following the procedure.</li>
	<li>House mice in pairs separated by cage dividers to prevent isolation whilst preventing fighting behaviors. Monitor mice at least daily until the end of the experiments (28 days) with close attention to their wellbeing and increased frequency of monitoring if there are any concerns.</li>
</ol>

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None.
FUNDING STATEMENT:
Christopher D. Roche was supported by a Sir John Loewenthal Scholarship 2019 (University of Sydney), the Le Gros Legacy Fund New Zealand (PhD012019) and a Heart Research Australia PhD Scholarship (2019-02). Carmine Gentile was supported by a University of Sydney Kick-Start Grant, University of Sydney Chancellor&#39;s Doctoral Incentive Programme Grant, UTS Seed Funding, Catholic Archdiocese of Sydney Grant for Adult Stem Cell Research and a Sydney Medical School Foudation Cardiothoracic Surgery Research Grant.

The method facilitates the operator to efficiently transplant a bioprinted patch by applying it to the epicardial surface of an infarcted mouse heart following permanent LAD ligation. In this feasibility-focused method, we are able to perform this procedure on eight mice per working day (including preparation of the room before and afterwards). A bioprinting run producing eight 1 cm2 patches in wells of six-well plates takes 2-3 hours (including preparation time before and after). We used the sterile inside of a surgical scalpel packet as the scoop for our patch, which is readily accessible and generally adds minimal cost, utilizing the natural adhesive properties of the alginate/gelatine hydrogel patch to drape the patch across the anterior infarcted surface of the heart. In our experience, the protocol for LAD ligation in mice is operator dependent and a lower mortality rate at 28 days can be achieved with experienced operators specialized in one model. Van den Borne et al.16 reported that C57BL6 mice present a 44% mortality following permanent LAD ligation at 28 days without the application of a patch, which is higher than the upper limit of 30% that we observed with the method.
The intubation step is critical and in-and-of-itself can be a source of mortality for mice unless performed by a skilled operator. It is made difficult due to the tiny size of the trachea, which is why magnifying glasses are worn by the operator for this step. We use injected ketamine/xylazine as well as inhaled isofluorane for induction of anesthetic so that the mouse is deeply anesthetized at relatively low doses of each drug. Therefore, there is no risk for the mouse to wake up during this intubation step but the high mortality associated with high single-drug doses is avoided. Atropine was also given to counteract side effects such as bradycardia and hypersalivation. The use of a spotlight applied to the throat externally lights up the trachea internally so it is more visible and the vocal cords must be visualized opening and closing with the mouse&#8217;s respiratory rate (usually ~120 breaths per minute). It is critical to position the mouse perfectly (which is why a hard surface is preferred rather than a warming mat underneath the mouse for this step) with the two incisor teeth held by a looped thread and the tongue retracted extremely gently with blunt forceps/pair of spatulas to open the mouth and visualize the trachea. Once the intubation is completed, the operator must be careful not to dislodge the tube in the transfer from the intubation area to the operating bed (which does have a heat mat underneath it to prevent hypothermia). When connecting the breathing tube to the ventilator apparatus, it is critical to stabilize the tube with one hand and connect the ventilator circuit with the other, so that there is minimal movement of the breathing tube such as pushing it more deeply into the trachea when connecting the ventilator segment of the tubing.
In this study, we used alginate 4% (w/v)/gelatine 8% (w/v) in Dulbecco&#8217;s Modified Eagle Medium (DMEM). Alginate/gelatine hydrogels are known for their biocompatibility, low cost and biomechanical properties making them useful for 3D tissue engineering strategies23. These hydrogels can be crosslinked by mild gelation by adding calcium ions, which allows for viscosity to be altered. After bioprinting, we applied calcium chloride (CaCl2) 2% (w/v) in phosphate-buffered saline (PBS) onto patches and then cultured them in DMEM in six well plates for 7-14 days before transplanting them. This was the optimal window after patches containing cardiac cells started to beat in culture but before patches started to disintegrate. Whilst CaCl2 could be added regularly throughout the post-bioprinting phase to reduce patch disintegration, we found that the intrinsic viscosity of the hydrogel was sufficient for patches to maintain their structure up to transplantation with only one initial dose of CaCl2.
The method allowed for successful transplantation without sutures (which may damage the heart) or glue (which may block the interface between the patch and the heart). Future studies may confirm the hypothesis that sutureless and glueless transplantation does not negatively impact engraftment in mice as it is critical that the patch does not slip off the heart or interfere with the lungs. Other studies assessing engraftment of patches in permanent LAD ligation models with patch-based repair3 have measured engrafted area (mm2) remaining with time24, the grafted patch thickness (&#956;m) remining with time25, quantification of transplanted cells by polymerase chain reaction (PCR)26 or bioluminescence photon emission flux of labelled live donor cells (a measure of photons emitted per second which can quantify labelled grafted cells surviving in living animals over time)27. Future studies may use these methods to further evaluate whether sutureless and glueless transplantation affects patch engraftment (as well as structural and functional effects on the host myocardium). Nonetheless, macroscopically after 28 days in vivo in our immunocompetent mice, the anterior mediastinum presented variable fibrinous material and adhesions. The mechanism of patch-based cardiac regeneration may be from stimulation of host macrophage inflammatory responses19 or secreted immunological factors20 rather than numerical cell replenishment. If inflammation plays a positive role, the presence of foreign hydrogel material may be beneficial. Alternatively, to reduce the presence of foreign material it may be beneficial if the hydrogel component disintegrates over time. In fact, some approaches use biomaterials which support cells initially and then disintegrate, leaving only tissue28,29. Future studies to fully analyze patch engraftment and better understand the mechanisms behind patch-based cardiac regeneration may lead to optimized experimental designs before translation to human trials2.
Overall, this protocol is likely to be widely feasible and also suited to testing multiple groups of 3D bioprinted patches, for instance with different cellular contents. Future directions for this method include the bioprinting of patches containing advanced hydrogels not previously tested in vivo or testing the effects of different autologous or allogenic stem cell-derived cells, for optimization before proceeding to large animal models.

A heart transplant is the gold standard treatment for patients with end-stage heart failure but there is a shortage of donor organs. It requires immune system suppression to prevent graft rejection and the one-year mortality rate is 15% worldwide1. Therefore, there is a longstanding incentive to regenerate the myocardium in preclinical animal models with a view to translating to human trials2,3,4,5,6,7,8,9. Recent advances in 3D bioprinting of stem cells or stem cell-derived cardiac cells have gained attention as a promising approach to regenerate the myocardium2,3,9,10,11,12.
The first human safety trials applying patches to regenerate the heart have been reported, with autologous bone marrow mononuclear cells suspended in collagen or embryonic stem cell-derived cardiac progenitor cells in fibrin, transplanted to the epicardium7,8,13. However, for a more precise, scalable, automatable and reproducible method, 3D bioprinting of optimized hydrogel patches to be applied to the epicardial surface of the heart is a promising approach to regenerate the myocardium for patients who would otherwise need a heart transplant2,10,11,12.
Before translation to human trials can occur, preclinical animal studies are needed. Preclinical in vivo models pursuing regeneration of the myocardium have been reported in pigs5, sheep14, rats6 and mice4. A common model of myocardial infarction (MI) in mice uses permanent ligation of the left anterior descending (LAD) coronary artery15,16. Among the different strains of mice used, permanent LAD ligation in C57BL6 mice has an acceptable survival rate and typically presents consistent remodeling and cardiac changes after MI16. In rodent models, several approaches have been described where cardiac tissue has been applied to the heart in pursuit of effective regeneration of damaged myocardium4,6,17. While large animals still represent a more clinically relevant model to test cardiac regenerative properties5,14, the versatility and feasibility of the mouse model lends itself to this fast-moving area of study. This may avoid some of the pitfalls typical of large animal studies, including (but not limited to): 1) high animal mortality (unless diagonal coronary arteries are ligated leading to unpredictable segmental infarcts14, or the distal end of the LAD is occluded followed by reperfusion instead of permanent ligation5); 2) ethical issues with the relatively increased harm caused by large animal protocols compared to mice18; 3) increased cost and/or feasibility issues, for instance the relative unavailability of large animal equipment such as MRI scanners14. It is also important to consider that given the extensive duration and commitment typical of large animal studies, they have the potential to become outdated before they are finished, especially with the rapid developments typical of this field. For instance, it is only recently that the critical role played by inflammatory cells and mediators in regulating cardiac regeneration has emerged19,20. Furthermore, the critical role of preclinical studies, such as small animal models, has been highlighted by a Lancet Commission as an essential step to gain robust knowledge before moving to human trials21.
To facilitate progress in understanding mechanisms and optimizing conditions for patch-based cardiac regeneration approaches in vivo, we present a novel approach describing a &#8216;scoop and drape&#8217; method to apply a 3D bioprinted alginate/gelatine hydrogel patch to the surface of infarcted hearts in C57BL6 mice. The aim of this approach is to provide a versatile in vivo model to test 3D bioprinted patches that are likely to be feasible in broad research contexts for the rapidly-evolving field of cardiac regeneration2. This method could be adapted to test patches generated by non-bioprinting methods, different hydrogels and autologous or allogenic stem cell-derived cells within patches in vivo. However, detailed consideration of bioprinting, hydrogels or cell types is beyond the scope of this study which focuses on the surgical transplantation method.
The advantages of the protocol include that the myocardial infarction and application of a bioprinted patch are performed in one surgical procedure that can be performed quickly, with readily-available, cost effective laboratory tools and with a relatively low mortality rate. It also typically allows for a higher number of animals than large animal models in a smaller space, which permits robust comparison of multiple experimental groups, particularly useful for multiple group comparison in vivo. On the other hand, this protocol has the disadvantages that: 1) the mouse model is more distant from human heart size, anatomy and physiology than in large animal models and it does not directly translate into humans; 2) the murine LAD branches proximally, with significant variability between individual mice, which leads to infarct size variability (a problem shared with large animal models); 3) the patch must be applied over the whole anterior heart surface, which is less precise than applying over a specific infarct area; and 4) the patch is applied immediately at the time of MI (for human use it is likely to be more clinically useful to develop a patch for application to the chronically infarcted failing heart months following the initial MI14).
Nonetheless, if chosen appropriately according to the hypothesis being tested, this protocol can provide critical in vivo data quickly, with high n numbers, in a way that is consistent with the materials, budget and expertise available in most laboratories. Compared to large animal models, it is an in vivo model that is versatile enough to adapt to emerging 3D bioprinting technologies (for example by the relative ease of performing pilot studies to test feasibility and safety before moving to larger animal models). It would be well-suited for researchers who want to generate in vivo data efficiently and inexpensively, perhaps running multiple comparisons of 3D bioprinted patches with different bioprinting parameters, cells or hydrogels in the patches. It would be especially useful for testing the interactions of different mixtures of stem cells and stem cell-derived cells with hydrogels in vivo without excess wastage of expensive cell lineages or other materials that might occur if using large scale patches. Using a mouse model would also facilitate testing of patches containing species-compatible mouse-derived cell and stem cell lineages or human-derived cells where uniform mice with a specific immune deficiency are desirable. Additionally, testing in genetically modified mouse strains could allow researchers to isolate the effects of specific genes on signaling pathways and in specific cell types relevant to cardiovascular disease, which would not currently be possible in a large animal model.

<table><tbody><tr><td>3-0 non-absorbable black braided treated silk</td><td>Ethicon</td><td>232G</td><td></td></tr><tr><td>6-0, 24&amp;rdquo; (60 cm) Prolene (polypropylene) suture, blue monofilament</td><td>Ethicon</td><td>8805H</td><td></td></tr><tr><td>7-0, 18&amp;rdquo; (45 cm) silk black braided</td><td>Ethicon</td><td>768G</td><td></td></tr><tr><td>Adjustable stereo microscope with 6.4x magnification</td><td>Olympus</td><td>SZ 3060 STU1</td><td></td></tr><tr><td>Anitisedan (atipamezole)</td><td>Zoetis</td><td>N/A</td><td></td></tr><tr><td>Atropine sulphate 0.6 mg, 1 mL vials, 10 pack</td><td>Symbion Pharmacy Services</td><td>ATRO S I2</td><td></td></tr><tr><td>Bupivacaine, 20 mL, 5 vials</td><td>Baxter Heathcare</td><td>BUPI I C01</td><td></td></tr><tr><td>Temvet (buprenorphine), 300 &amp;micro;g/mL, 10 mL bottle</td><td>Troy Laboratories</td><td>TEMV I 10</td><td></td></tr><tr><td>Curved-tip forceps</td><td>Kent Scientific</td><td>INS650915-4</td><td>Iris dressing forceps, 10 cm-long curved dressing forceps; 0.8 mm serrated tips; stainless steel.</td></tr><tr><td>Dissecting scissors for cutting muscle/skin</td><td>Kent Scientific</td><td>INS600393-G</td><td>Dissecting scissors, straight, 10 cm long</td></tr><tr><td>Endotracheal intubation kit</td><td>Kent Scientific</td><td>ETI-MSE</td><td>Including intubation catheter/tube (20 G), fibre-optic light source and dental spatula</td></tr><tr><td>Fine scissors</td><td>Kent Scientific</td><td>INS600124</td><td>McPherson-Vannas micro scissors, 8 cm long, straight, 0.1 mm tips, 5 mm blades; stainless steel.</td></tr><tr><td>Lasix (furosemide) 20 mg, 2 mL, 5 pack</td><td>Sigma Company</td><td>LASI A 1</td><td></td></tr><tr><td>Heat pad for animal recovery post-op</td><td>Passwell</td><td>PAD</td><td>Passwell Cosy Heat Pad for Animals - 26cm x 36cm; 10 Watts; Soft PVC Cover</td></tr><tr><td>Ketamine 100 mg, 50 mL</td><td>CEVA Animal Heath</td><td>KETA I 1</td><td></td></tr><tr><td>Needle holder</td><td>Kent Scientific</td><td>INS600137</td><td>Castroviejo needle holder, serrated, 14 cm long, 1.2 mm jaws with lock</td></tr><tr><td>PhysioSuite with MouseVent G500 automatic ventilator</td><td>Kent Scientific</td><td>PS-MVG</td><td></td></tr><tr><td>Puralube Vet Opthalmic Ointment (sterile occular lubricant)</td><td>Dechra</td><td>17033-211-38</td><td></td></tr><tr><td>Self-retaining toothed mouse retractor</td><td>Kent Scientific</td><td>INS600240</td><td>ALM serrated self-retaining retractor, 7 cm long</td></tr><tr><td>Straight forceps</td><td>Kent Scientific</td><td>INS650908-4</td><td>Super fine dressing forceps, 12.5 cm Long, serrated tips, 0.35 x 0.10 mm; stainless steel.</td></tr><tr><td>Surgical magnifying glasses</td><td>Kent Scientific</td><td>SL-001</td><td></td></tr><tr><td>VetFlo vaporizer</td><td>Kent Scientific</td><td>VetFlo-1205S-M</td><td></td></tr><tr><td>Xylazine 100 mg, 50 mL</td><td>Randlab</td><td>XYLA I R01</td><td></td></tr></tbody></table>

transplantation of a 3d bioprinted patch in a murine model of myocardial infarction

Testing regenerative properties of 3D bioprinted cardiac patches in vivo using murine models of heart failure via permanent left anterior descending (LAD) ligation is a challenging procedure and has a high mortality rate due to its nature. We developed a method to consistently transplant bioprinted patches of cells and hydrogels onto the epicardium of an infarcted mouse heart to test their regenerative properties in a robust and feasible way. First, a deeply anesthetized mouse is carefully intubated and ventilated. Following left lateral thoracotomy (surgical opening of the chest), the exposed LAD is permanently ligated and the bioprinted patch transplanted onto the epicardium. The mouse quickly recovers from the procedure after chest closure. The advantages of this robust and quick approach include a predicted 28-day mortality rate of up to 30% (lower than the 44% reported by other studies using a similar model of permanent LAD ligation in mice). Moreover, the approach described in this protocol is versatile and could be adapted to test bioprinted patches using different cell types or hydrogels where high numbers of animals are needed to optimally power studies. Overall, we present this as an advantageous approach which may change preclinical testing in future studies for the field of cardiac regeneration and tissue engineering.

At transplantation, the viscosity of the patch at room temperature (without additional crosslinker being applied) allowed it to &#8216;drape&#8217; over the contours of the heart (Figure 1) and move dynamically with the cardiac cycle. After the surgery, we left the patches for 28 days in vivo as studies have found this to be a suitable time period allowing for patch effects on host cardiac function3,4 (although it has been reported that full functional effects may not be seen until three months after transplantation)22. The photograph of a patch shown in situ on a mouse heart in Figure 1 was taken immediately after application, showing the ability of the patch to drape over the heart at transplantation. This representative result shows that the hydrogel allows for the patch to mold to the contours of the heart and where excessive tension occurred the hydrogel was able to split as shown by the bare (hydrogel-free) triangular area in Figure 1 (indicated by a black star in the image). Survival data (Kaplan-Meier survival curves) are shown in Figure 2 compared to mice undergoing a sham procedure (passage of a needle and suture under the LAD without ligation followed by closure of the mouse chest).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61675/61675fig01.jpg" /> 
Figure 1: A bioprinted cardiac patch applied onto the epicardium of a C57BL6 mouse heart.&#160;A 10 mm x 10 mm x 0.4 mm bioprinted patch (immediately after transplantation) containing hydrogel (alginate 4% (w/v)/gelatine 8% (w/v) in media) is shown draped over the infarcted area and adhering to the epicardial surface (white arrowheads and dotted lines = border of the patch). The patch viscosity allows it to mold to the contours of the heart and where excessive tension occurred at the superior aspect the patch has split to make a triangular bare area not covered by hydrogel (black star). <a href="https://www.jove.com/files/ftp_upload/61675/61675fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61675/61675fig02.jpg" /> 
Figure 2: Kaplan-Meier survival analysis through 28 days post-MI.&#160;Nine mice in the procedural group died (n=38) to give an overall mortality rate of 24%. <a href="https://www.jove.com/files/ftp_upload/61675/61675fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Video still image (video time point 01:54 &#8211; 01:55) showing the left auricle (left atrial appendage). The arrow points to the inferomedial tip of the left auricle which is visible as a triangular structure at the superior left edge of the heart. In case the LAD is not clearly visualized, the tip of the left auricle can be used as a landmark for needle entry to pass a suture under the LAD. The entry point is 1 mm inferior and medial to the inferiormost point the tip of the left auricle reaches during dynamic movements of the heart (black arrow shows inferomedial tip of the left auricle). <a href="https://cloudflare.jove.com/files/ftp_upload/61675/61675supfig01.jpg" target="_blank">Please click here to download this figure.</a>

Watch this Scientific Journal Video about Transplantation of a 3D Bioprinted Patch in a Murine Model of Myocardial Infarction at JoVE.com

Transplantation of a 3D Bioprinted Patch in a Murine Model of Myocardial Infarction

This protocol aims to transplant a 3D bioprinted patch onto the epicardium of infarcted mice modeling heart failure. It includes details regarding anesthesia, the surgical chest opening, permanent ligation of the left anterior descending (LAD) coronary artery and application of a bioprinted patch onto the infarcted area of the heart.

Testing regenerative properties of 3D bioprinted cardiac patches in vivo using murine models of heart failure via permanent left anterior descending (LAD) ...

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Research

JoVE Journal

Bioengineering

6.5K Views.  The University of Sydney. This protocol aims to transplant a 3D bioprinted patch onto the epicardium of infarcted mice modeling heart failure. It includes details regarding anesthesia, the surgical chest opening, permanent ligation of the left anterior descending (LAD) coronary artery and application of a bioprinted patch onto the infarcted area of the heart.

Video: Transplantation of a 3D Bioprinted Patch in a Murine Model of Myocardial Infarction

Generation of Aligned Functional Myocardial Tissue Through Microcontact Printing

Advanced heart failure represents a major unmet clinical challenge, arising from the loss of viable and/or fully functional cardiac muscle cells. Despite optimum drug therapy, heart failure represents a leading cause of mortality and morbidity in the developed world. A major challenge in drug development is the identification of cellular assays that accurately recapitulate normal and diseased human myocardial physiology in vitro. Likewise, the major challenges in regenerative cardiac biology revolve around the identification and isolation of patient-specific cardiac progenitors in clinically relevant quantities. These cells have to then be assembled into functional tissue that resembles the native heart tissue architecture. Microcontact printing allows for the creation of precise micropatterned protein shapes that resemble structural organization of the heart, thus providing geometric cues to control cell adhesion spatially. Herein we describe our approach for the isolation of highly purified myocardial cells from pluripotent stem cells differentiating in vitro, the generation of cell growth surfaces micropatterned with extracellular matrix proteins, and the assembly of the stem cell-derived cardiac muscle cells into anisotropic myocardial tissue.

The generation of aligned myocardial tissue is a key requirement for adapting the recent advances in stem cell biology to clinically useful purposes. Herein we describe a microcontact printing approach for the precise control of cell shape and function. Using highly purified populations of embryonic stem cell derived cardiac progenitors, we then generate anisotropic functional myocardial tissue.

Advanced heart failure represents a major unmet clinical challenge,  arising from the loss of viable and/or fully functional cardiac muscle  cells. ...

Intramyocardial Cell Delivery: Observations in Murine Hearts

Previous studies showed that cell delivery promotes cardiac function amelioration by release of cytokines and factors that increase cardiac tissue revascularization and cell survival. In addition, further observations revealed that specific stem cells, such as cardiac stem cells, mesenchymal stem cells and cardiospheres have the ability to integrate within the surrounding myocardium by differentiating into cardiomyocytes, smooth muscle cells and endothelial cells.
Here, we present the materials and methods to reliably deliver noncontractile cells into the left ventricular wall of immunodepleted mice. The salient steps of this microsurgical procedure involve anesthesia and analgesia injection, intratracheal intubation, incision to open the chest and expose the heart and delivery of cells by a sterile 30-gauge needle and a precision microliter syringe.
Tissue processing consisting of heart harvesting, embedding, sectioning and histological staining showed that intramyocardial cell injection produced a small damage in the epicardial area, as well as in the ventricular wall. Noncontractile cells were retained into the myocardial wall of immunocompromised mice and were surrounded by a layer of fibrotic tissue, likely to protect from cardiac pressure and mechanical load.

Intramyocardial cell delivery in murine models of cardiovascular diseases, such as hypertension or myocardial infarction, is widely used to test the therapeutic potential of different cell types in regenerative studies. Therefore, a detailed description and a clear visualization of this surgical procedure will help to define the limits and advantages of cardiovascular cell therapeutic analyses in small rodents.

Previous studies showed that cell delivery promotes cardiac function amelioration by release of cytokines and factors that increase cardiac tissue ...

Medicine

LAD-Ligation: A Murine Model of Myocardial Infarction

Research models of infarction and myocardial ischemia are essential to investigate the acute and chronic pathobiological and pathophysiological processes in myocardial ischemia and to develop and optimize future treatment.
Two different methods of creating myocardial ischemia are performed in laboratory rodents. The first method is to create cryo infarction, a fast but inaccurate technique, where a cryo-pen is applied on the surface of the heart (1-3). Using this method the scientist can not guarantee that the cryo-scar leads to ischemia, also a vast myocardial injury is created that shows pathophysiological side effects that are not related to myocardial infarction. The second method is the permanent ligation of the left anterior descending artery (LAD). Here the LAD is ligated with one single stitch, forming an ischemia that can be seen almost immediately. By closing the LAD, no further blood flow is permitted in that area, while the surrounding myocardial tissue is nearly not affected. This surgical procedure imitates the pathobiological and pathophysiological aspects occurring in infarction-related myocardial ischemia.
The method introduced in this video demonstrates the surgical procedure of a mouse infarction model by ligating the LAD. This model is convenient for pathobiological and pathophysiological as well as immunobiological studies on cardiac infarction. The shown technique provides high accuracy and correlates well with histological sections.

This video demonstrates how to use a fast and reliable model to study pathobiological and pathophysiological processes of myocardial ischemia.

Research models of infarction and myocardial ischemia are essential to investigate the acute and chronic pathobiological and pathophysiological processes ...

Biology

Creation of Cardiac Tissue Exhibiting Mechanical Integration of Spheroids Using 3D Bioprinting

This protocol describes 3D bioprinting of cardiac tissue without the use of biomaterials, using only cells. Cardiomyocytes, endothelial cells and fibroblasts are first isolated, counted and mixed at desired cell ratios. They are co-cultured in individual wells in ultra-low attachment 96-well plates. Within 3 days, beating spheroids form. These spheroids are then picked up by a nozzle using vacuum suction and assembled on a needle array using a 3D bioprinter. The spheroids are then allowed to fuse on the needle array. Three days after 3D bioprinting, the spheroids are removed as an intact patch, which is already spontaneously beating. 3D bioprinted cardiac patches exhibit mechanical integration of component spheroids and are highly promising in cardiac tissue regeneration and as 3D models of heart disease.

This protocol describes 3D bioprinting of cardiac tissue without the use of biomaterials. 3D bioprinted cardiac patches exhibit mechanical integration of component spheroids and are highly promising in cardiac tissue regeneration and as 3D models of heart disease.

This protocol describes 3D bioprinting of cardiac tissue without the use of biomaterials, using only cells. Cardiomyocytes, endothelial cells and ...

A Hydrogel Construct and Fibrin-based Glue Approach to Deliver Therapeutics in a Murine Myocardial Infarction Model.

The murine MI model is widely recognized in the field of cardiovascular disease, and has consistently been used as a first step to test the efficacy of treatments in vivo1. The traditional, established protocol has been further fine-tuned to minimize the damage to the animal. Notably, the pectoral muscle layers are teased away rather than simply cut, and the thoracotomy is approached intercostally as opposed to breaking the ribs in a sternotomy, preserving the integrity of the ribcage. With these changes, the overall stress on the animal is decreased. 
Stem cell therapies aimed to alleviate the damage caused by MIs have shown promise over the years for their pro-angiogenic and anti-apoptotic benefits. Current approaches of delivering cells to the heart surface typically involve the injection of the cells either near the damaged site, within a coronary artery, or into the peripheral blood stream2-4. While the cells have proven to home to the damaged myocardium, functionality is limited by their poor engraftment at the site of injury, resulting in diffusion into the blood stream5. This manuscript highlights a procedure that overcomes this obstacle with the use of a cell-encapsulated hydrogel patch. The patch is fabricated prior to the surgical procedure and is placed on the injured myocardium immediately following the occlusion of the left coronary artery. To adhere the patch in place, biocompatible external fibrin glue is placed directly on top of the patch, allowing for it to dry to both the patch and the heart surface. This approach provides a novel adhesion method for the application of a delicate cell-encapsulating therapeutic construct.

This protocol aims to alleviate the limitation of poor cell engraftment for stem cell treatment of myocardial infarctions through the use of a hydrogel system and a fibrin-based glue. With this approach, cell-to-tissue contact post-infarction can be maintained, increasing the therapeutic potential of beneficial agents at the site of injury.

The murine MI model is widely recognized in the field of cardiovascular disease, and has consistently been used as a first step to test the efficacy of ...

3D Whole-heart Myocardial Tissue Analysis

Cardiac regenerative therapies aim to protect and repair the injured heart in patients with ischemic heart disease. By injecting stem cells or other biologicals that enhance angio- or vasculogenesis into the infarct border zone (IBZ), tissue perfusion is improved, and the myocardium can be protected from further damage. For maximum therapeutic effect, it is hypothesized that the regenerative substance is best delivered to the IBZ. This requires accurate injections and has led to the development of new injection techniques. To validate these new techniques, we have designed a validation protocol based on myocardial tissue analysis. This protocol includes whole-heart myocardial tissue processing that enables detailed two-dimensional (2D) and three-dimensional (3D) analysis of the cardiac anatomy and intramyocardial injections. In a pig, myocardial infarction was created by a 90-min occlusion of the left anterior descending coronary artery. Four weeks later, a mixture of a hydrogel with superparamagnetic iron oxide particles (SPIOs) and fluorescent beads was injected in the IBZ using a minimally-invasive endocardial approach. 1 h after the injection procedure, the pig was euthanized, and the heart was excised and embedded in agarose (agar). After the solidification of the agar, magnetic resonance imaging (MRI), slicing of the heart, and fluorescence imaging were performed. After image post-processing, 3D analysis was performed to assess the IBZ targeting accuracy. This protocol provides a structured and reproducible method for the assessment of the targeting accuracy of intramyocardial injections into the IBZ. The protocol can be easily used when the processing of scar tissue and/or validation of the injection accuracy of the whole heart is desired.

This protocol describes a novel method for the 3D comparison of whole-heart myocardial tissue with MRI. This is designed for the accurate assessment of intramyocardial injections in the infarct border zone of a chronic porcine model of myocardial infarction.

Cardiac regenerative therapies aim to protect and repair the injured heart in patients with ischemic heart disease. By injecting stem cells or other ...

Implantation of hiPSC-derived Cardiac-muscle Patches after Myocardial Injury in a Guinea Pig Model

Due to the limited regeneration capacity of the heart in adult mammals, myocardial infarction results in an irreversible loss of cardiomyocytes. This loss of relevant amounts of heart muscle mass can lead to the heart failure. Besides heart transplantation, there is no curative treatment option for the end-stage heart failure. In times of organ donor shortage, organ independent treatment modalities are needed. Left-ventricular assist devices are a promising therapy option, however, especially as destination therapy, limited by its side-effects like stroke, infections and bleedings. In recent years, several cardiac repair strategies including stem cell injection, cardiac progenitors or myocardial tissue engineering have been investigated. Recent improvements in cell biology allow for the differentiation of large amounts of cardiomyocytes derived from human induced pluripotent stem cells (iPSC). One of the cardiac repair strategies currently under evaluation is to transplant artificial heart tissue. Engineered heart tissue (EHT) is a three-dimensional in vitro created cardiomyocyte network, with functional properties of native heart tissue. We have created EHT-patches from hiPSC derived cardiomyocytes. Here we present a protocol for the induction of left ventricular myocardial cryoinjury in a guinea pig, followed by implantation of hiPSC derived EHT on the left ventricular wall.

Here we present a protocol for the induction of left ventricular cryoinjury followed by the implantation of a cardiac muscle patch, derived from human iPS-cell cardiomyocytes in a guinea pig model.

Due to the limited regeneration capacity of the heart in adult mammals, myocardial infarction results in an irreversible loss of cardiomyocytes. This loss ...

Murine Myocardial Infarction Model using Permanent Ligation of Left Anterior Descending Coronary Artery

Myocardial infarction (MI) and acute coronary diseases are among the most prominent causes of death in population with western lifestyle. The murine models of MI with permanent ligation of left-anterior descending (LAD) coronary artery closely mimics MI in humans. Murine models benefit from the extensive genetic engineering available nowadays. Here we propose a reproducible murine surgical model of myocardial infarction by permanent LAD coronary ligation. Our technique comprises anesthesia with ketamine/xylazine that can be rapidly reversed by administration of an antagonist, intubation without tracheotomy for mechanical-assisted ventilation, ventilation with application of extrinsic positive end-expiratory pressure (PEEP) to avoid alveolar collapse, a thoracotomy method limiting to the minimum surgical lesions made to skeletal muscles, and lung inflation without thoracentesis. This method is sparsely invasive, reproducible and reduces post-surgery mortality and complications.

Herein we describe a surgical procedure showing how to achieve permanent ligation of the left-anterior descending coronary artery in mice. This model is of high relevance to investigate the pathophysiology of myocardial infarction and the concomitant biological processes.

Myocardial infarction (MI) and acute coronary diseases are among the most prominent causes of death in population with western lifestyle. The murine ...

Mouse Heterotopic Cervical Cardiac Transplantation Utilizing Vascular Cuffs

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after transplantation, and the impact of immunomodulatory therapies on graft rejection. Heterotopic cervical heart transplantation in mice was first described in 1991 using sutured anastomoses and subsequently modified to include cuff techniques. This modification allowed for improved success rates, and since then, there have been multiple reports that have proposed further technical improvements. However, translation into more widespread utilization remains limited due to the technical difficulty associated with graft anastomoses, which requires precision to achieve adequate length and caliber of the cuffs to avoid vascular anastomotic twisting or excessive tension, which can result in damage to the graft. The present protocol describes a modified technique for performing heterotopic cervical cardiac transplantation in mice which involves cuff placement on the recipient's common carotid artery and the donor's pulmonary artery in alignment with the direction of the blood flow.

Mouse cardiac transplantation models represent valuable research tools for studying transplantation immunology. The present protocol details mouse heterotopic cervical cardiac transplantation that involves the placement of cuffs on the recipient's common carotid artery and the donor's pulmonary artery trunk to allow for laminar blood flow.

Murine models of cardiac transplantation are frequently utilized to study ischemia-reperfusion injury, innate and adaptive immune responses after ...

Immunology and Infection

Intramyocardial Transplantation of MSC-Loading Injectable Hydrogels after Myocardial Infarction in a Murine Model

One of the major issues facing current cardiac stem cell therapies for preventing postinfarct heart failure is the low retention and survival rates of transplanted cells within the injured myocardium, limiting their therapeutic efficacy. Recently, the use of scaffolding biomaterials has gained attention for improving and maximizing stem cell therapy. The objective of this protocol is to introduce a simple and straightforward technique to transplant bone marrow-derived mesenchymal stem cells (MSCs) using injectable hydroxyphenyl propionic acid (GH) hydrogels; the hydrogels are favorable as a cell delivery platform for cardiac tissue engineering applications due to their ability to be cross-linked in situ and high biocompatibility. We present a simple method to fabricate MSC-loading GH hydrogels (MSC/hydrogels) and evaluate their survival and proliferation in three-dimensional (3D) in vitro culture. In addition, we demonstrate a technique for intramyocardial transplantation of MSC/hydrogels in mice, describing a surgical procedure to induce myocardial infarction (MI) via left anterior descending (LAD) coronary artery ligation and subsequent MSC/hydrogels transplantation.

Stem cell-based therapy has emerged as an efficient strategy to repair injured cardiac tissues after myocardial infarction. We provide an optimal in vivo application for stem cell transplantation using gelatin hydrogels that are able to be enzymatically cross-linked.

One of the major issues facing current cardiac stem cell therapies for preventing postinfarct heart failure is the low retention and survival rates of ...