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&#8221; (60 cm) Prolene (polypropylene) suture, blue monofilament</td>
			<td>Ethicon</td>
			<td>8805H</td>
			<td></td>
		</tr>
		<tr>
			<td>7-0, 18&#8221; (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 suplhate 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 &#181;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

Tissue Engineering of the Intestine in a Murine Model

Tissue-engineered small intestine (TESI) has successfully been used to rescue Lewis rats after massive small bowel resection, resulting in return to preoperative weights within 40 days.1 In humans, massive small bowel resection can result in short bowel syndrome, a functional malabsorptive state that confers significant morbidity, mortality, and healthcare costs including parenteral nutrition dependence, liver failure and cirrhosis, and the need for multivisceral organ transplantation.2 In this paper, we describe and document our protocol for creating tissue-engineered intestine in a mouse model with a multicellular organoid units-on-scaffold approach. Organoid units are multicellular aggregates derived from the intestine that contain both mucosal and mesenchymal elements,3 the relationship between which preserves the intestinal stem cell niche.4 In ongoing and future research, the transition of our technique into the mouse will allow for investigation of the processes involved during TESI formation by utilizing the transgenic tools available in this species.5The availability of immunocompromised mouse strains will also permit us to apply the technique to human intestinal tissue and optimize the formation of human TESI as a mouse xenograft before its transition into humans. Our method employs good manufacturing practice (GMP) reagents and materials that have already been approved for use in human patients, and therefore offers a significant advantage over approaches that rely upon decellularized animal tissues. The ultimate goal of this method is its translation to humans as a regenerative medicine therapeutic strategy for short bowel syndrome.

This article and the accompanying video present our protocol for generating tissue-engineered intestine in the mouse, using an organoid units-on-scaffold approach.

Tissue-engineered small intestine (TESI) has successfully been used to rescue Lewis rats after massive small bowel resection, resulting in return to ...

Cell-based Therapy for Heart Failure in Rat: Double Thoracotomy for Myocardial Infarction and Epicardial Implantation of Cells and Biomatrix

Cardiac cell therapy has gained increasing interest and implantation of biomaterials associated with cells has become a major issue to optimize myocardial cell delivery. Rodent model of myocardial infarction (MI) consisting of Left Anterior Descending Artery (LAD) ligation has commonly been performed via a thoracotomy; a second open-heart surgery via a sternotomy has traditionally been performed for epicardial application of the treatment. Since the description of LAD ligation model, post-surgery mortality rate has dropped from 35-13%, however the second surgery has remained critical. In order to improve post-surgery recovery and reduce pain and infection, minimally invasive surgical procedures are presented.&#160;Two thoracotomies were performed, the initial one for LAD ligation and the second one for treatment epicardial administration. Biografts consisting of cells associated with solid or gel type matrices were applied onto the infarcted area.&#160;LAD ligation resulted in loss of heart function as confirmed by echocardiography performed after 2 and 6 weeks. Goldner trichrome staining performed on heart sections confirmed transmural scar formation. First and second surgeries resulted in less that 10%&#160;post-operative mortality.&#160;

Implantation of a biograft to treat myocardial infarction induced by LAD ligation in a rodent model has conventionally required two open-heart surgeries. In order to reduce mortality and provide optimal conditions for fixation of solid and gelatinous biomatrices associated with cells, minimally invasive procedures have been developed.

Cardiac cell therapy has gained increasing interest and implantation of biomaterials associated with cells has become a major issue to optimize myocardial ...

Production, Characterization and Potential Uses of a 3D Tissue-engineered Human Esophageal Mucosal Model

The incidence of both esophageal adenocarcinoma and its precursor, Barrett&#8217;s Metaplasia, are rising rapidly in the western world. Furthermore esophageal adenocarcinoma generally has a poor prognosis, with little improvement in survival rates in recent years. These are difficult conditions to study and there has been a lack of suitable experimental platforms to investigate disorders of the esophageal mucosa.
A model of the human esophageal mucosa has been developed in the MacNeil laboratory which, unlike conventional 2D cell culture systems, recapitulates the cell-cell and cell-matrix interactions present in vivo and produces a mature, stratified epithelium similar to that of the normal human esophagus. Briefly, the model utilizes non-transformed normal primary human esophageal fibroblasts and epithelial cells grown within a porcine-derived acellular esophageal scaffold. Immunohistochemical characterization of this model by CK4, CK14, Ki67 and involucrin staining demonstrates appropriate recapitulation of the histology of the normal human esophageal mucosa.
This model provides a robust, biologically relevant experimental model of the human esophageal mucosa. It can easily be manipulated to investigate a number of research questions including the effectiveness of pharmacological agents and the impact of exposure to environmental factors such as alcohol, toxins, high temperature or gastro-esophageal refluxate components. The model also facilitates extended culture periods not achievable with conventional 2D cell culture, enabling, inter alia, the study of the impact of repeated exposure of a mature epithelium to the agent of interest for up to 20 days. Furthermore, a variety of cell lines, such as those derived from esophageal tumors or Barrett&#8217;s Metaplasia, can be incorporated into the model to investigate processes such as tumor invasion and drug responsiveness in a more biologically relevant environment.

This manuscript describes the production, characterization and potential uses of a tissue engineered 3D esophageal construct prepared from normal primary human esophageal fibroblast and squamous epithelial cells seeded within a de-cellularized porcine scaffold. The results demonstrate the formation of a mature stratified epithelium similar to the normal human esophagus.

The incidence of both esophageal adenocarcinoma and its precursor, Barrett&#8217;s Metaplasia, are rising rapidly in the western world. Furthermore ...

A Hormone-responsive 3D Culture Model of the Human Mammary Gland Epithelium

The process of mammary epithelial morphogenesis is influenced by hormones. The study of hormone action on the breast epithelium using 2D cultures is limited to cell proliferation and gene expression endpoints. However, in the organism, mammary morphogenesis occurs in a 3D environment. 3D culture systems help bridge the gap between monolayer cell culture (2D) and the complexity of the organism. Herein, we describe a 3D culture model of the human breast epithelium that is suitable to study hormone action. It uses the commercially available hormone-responsive human breast epithelial cell line, T47D, and rat tail collagen type 1 as a matrix. This 3D culture model responds to the main mammotropic hormones: estradiol, progestins and prolactin. The influence of these hormones on epithelial morphogenesis can be observed after 1- or 2-week treatment according to the endpoint. The 3D cultures can be harvested for analysis of epithelial morphogenesis, cell proliferation and gene expression.

We describe a 3D culture model of the human breast epithelium that is suitable to study hormone action.

The process of mammary epithelial morphogenesis is influenced by hormones. The study of hormone action on the breast epithelium using 2D cultures is ...

Viability of Bioprinted Cellular Constructs Using a Three Dispenser Cartesian Printer

Tissue engineering has centralized its focus on the construction of replacements for non-functional or damaged tissue. The utilization of three-dimensional bioprinting in tissue engineering has generated new methods for the printing of cells and matrix to fabricate biomimetic tissue constructs. The solid freeform fabrication (SFF) method developed for three-dimensional bioprinting uses an additive manufacturing approach by depositing droplets of cells and hydrogels in a layer-by-layer fashion. Bioprinting fabrication is dependent on the specific placement of biological materials into three-dimensional architectures, and the printed constructs should closely mimic the complex organization of cells and extracellular matrices in native tissue. This paper highlights the use of the Palmetto Printer, a Cartesian bioprinter, as well as the process of producing spatially organized, viable constructs while simultaneously allowing control of environmental factors. This methodology utilizes computer-aided design and computer-aided manufacturing to produce these specific and complex geometries. Finally, this approach allows for the reproducible production of fabricated constructs optimized by controllable printing parameters.

A Cartesian bioprinter was designed and fabricated to allow multi-material deposition in precise, reproducible geometries, while also allowing control of environmental factors. Utilizing the three-dimensional bioprinter, complex and viable constructs may be printed and easily reproduced.

Tissue engineering has centralized its focus on the construction of replacements for non-functional or damaged tissue. The utilization of ...

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Historically, most cellular processes have been studied in only 2 dimensions. While these studies have been informative about general cell signaling mechanisms, they neglect important cellular cues received from the structural and mechanical properties of the local microenvironment and extracellular matrix (ECM). To understand how cells interact within a physiological ECM, it is important to study them in the context of 3 dimensional assays. Cell migration, cell differentiation, and cell proliferation are only a few processes that have been shown to be impacted by local changes in the mechanical properties of a 3-dimensional ECM. Collagen I, a core fibrillar component of the ECM, is more than a simple structural element of a tissue. Under normal conditions, mechanical cues from the collagen network direct morphogenesis and maintain cellular structures. In diseased microenvironments, such as the tumor microenvironment, the collagen network is often dramatically remodeled, demonstrating altered composition, enhanced deposition and altered fiber organization. In breast cancer, the degree of fiber alignment is important, as an increase in aligned fibers perpendicular to the tumor boundary has been correlated to poorer patient prognosis1. Aligned collagen matrices result in increased dissemination of tumor cells via persistent migration2,3. The following is a simple protocol for embedding cells within a 3-dimensional, fibrillar collagen hydrogel. This protocol is readily adaptable to many platforms, and can reproducibly generate both aligned and random collagen matrices for investigation of cell migration, cell division, and other cellular processes in a tunable, 3-dimensional, physiological microenvironment.

Preparation of 3D Collagen Gels and Microchannels for the Study of 3D Interactions In Vivo

Collagen is a core component of the ECM, and provides essential cues for several cellular processes ranging from migration to differentiation and proliferation. Provided here is a protocol for embedding cells within 3D collagen hydrogels, and a more advanced technique for generating randomized or aligned collagen matrices using PDMS microchannels.

Historically, most cellular processes have been studied in only 2 dimensions.  While these studies have been informative about general cell signaling ...

Micropatterning and Assembly of 3D Microvessels

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass based substrates, and hydrogel-based tube formation assays. These assays, while informative, do not recapitulate lumen geometry, proper extracellular matrix, and multi-cellular proximity, which play key roles in modulating vascular function. This manuscript describes an injection molding method to generate engineered vessels with diameters on the order of 100 &#181;m. Microvessels are fabricated by seeding endothelial cells in a microfluidic channel embedded within a native type I collagen hydrogel. By incorporating parenchymal cells within the collagen matrix prior to channel formation, specific tissue microenvironments can be modeled and studied. Additional modulations of hydrodynamic properties and media composition allow for control of complex vascular function within the desired microenvironment. This platform allows for the study of perivascular cell recruitment, blood-endothelium interactions, flow response, and tissue-microvascular interactions. Engineered microvessels offer the ability to isolate the influence from individual components of a vascular niche and precisely control its chemical, mechanical, and biological properties to study vascular biology in both health and disease.

This manuscript presents an injection molding method to engineer microvessels that recapitulate physiological properties of endothelium. The microfluidic-based process creates patent 3D vascular networks with tailorable conditions, such as flow, cellular composition, geometry, and biochemical gradients. The fabrication process and examples of potential applications are described.

In vitro platforms to study endothelial cells and vascular biology are largely limited to 2D endothelial cell culture, flow chambers with polymer or glass ...

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and dampen loads in the spine. The IVD consists of a proteoglycan-rich nucleus pulposus (NP) and a collagen-rich annulus fibrosis (AF) sandwiched by cartilaginous end-plates. Together with the adjacent vertebrae, the vertebrae-IVD structure forms a functional spine unit (FSU). These microstructures contain unique cell types as well as unique extracellular matrices. Whole organ culture of the FSU preserves the native extracellular matrix, cell differentiation phenotypes, and cellular-matrix interactions. Thus, organ culture techniques are particularly useful for investigating the complex biological mechanisms of the IVD. Here, we describe a high-throughput approach for culturing whole lumbar mouse FSUs that provides an ideal platform for studying disease mechanisms and therapies for the IVD. Furthermore, we describe several applications that utilize this organ culture method to conduct further studies including contrast-enhanced microCT imaging and three-dimensional high-resolution finite element modeling of the IVD.

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Whole organ culture of the intervertebral disc (IVD) preserves the native extracellular matrix, cell phenotypes, and cellular-matrix interactions. Here we describe an IVD culture system using mouse lumbar and caudal IVDs in their functional spinal units and several applications utilizing this system.

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and ...

Production of Elastin-like Protein Hydrogels for Encapsulation and Immunostaining of Cells in 3D

Two-dimensional (2D) tissue culture techniques have been essential for our understanding of fundamental cell biology. However, traditional 2D tissue culture systems lack a three-dimensional (3D) matrix, resulting in a significant disconnect between results collected in vitro and in vivo. To address this limitation, researchers have engineered 3D hydrogel tissue culture platforms that can mimic the biochemical and biophysical properties of the in vivo cell microenvironment. This research has motivated the need to develop material platforms that support 3D cell encapsulation and downstream biochemical assays. Recombinant protein engineering offers a unique toolset for 3D hydrogel material design and development by allowing for the specific control of protein sequence and therefore, by extension, the potential mechanical and biochemical properties of the resultant matrix. Here, we present a protocol for the expression of recombinantly-derived elastin-like protein (ELP), which can be used to form hydrogels with independently tunable mechanical properties and cell-adhesive ligand concentration. We further present a methodology for cell encapsulation within ELP hydrogels and subsequent immunofluorescent staining of embedded cells for downstream analysis and quantification.

Recombinant protein-engineered hydrogels are advantageous for 3D cell culture as they allow for complete tunability of the polymer backbone and therefore, the cell microenvironment. Here, we describe the process of recombinant elastin-like protein purification and its application in 3D hydrogel cell encapsulation.

Two-dimensional (2D) tissue culture techniques have been essential for our understanding of fundamental cell biology. However, traditional 2D tissue ...

A Novel Surgical Technique As a Foundation for In Vivo Partial Liver Engineering in Rat

Organ engineering is a novel strategy to generate liver organ substitutes that can potentially be used in transplantation. Recently, in vivo liver engineering, including in vivo organ decellularization followed by repopulation, has emerged as a promising approach over ex vivo liver engineering. However, postoperative survival was not achieved. The aim of this study is to develop a novel surgical technique of in vivo selective liver lobe perfusion in rats as a prerequisite for in vivo liver engineering. We generate a circuit bypass only through the left lateral lobe. Then, the left lateral lobe is perfused with heparinized saline. The experiment is performed with 4 groups (n = 3 rats per group) based on different perfusion times of 20 min, 2 h, 3 h, and 4 h. Survival, as well as the macroscopically visible change of color and the histologically determined absence of blood cells in the portal triad and the sinusoids, is taken as an indicator for a successful model establishment. After selective perfusion of the left lateral lobe, we observe that the left lateral lobe, indeed, turned from red to faint yellow. In a histological assessment, no blood cells are visible in the branch of the portal vein, the central vein, and the sinusoids. The left lateral lobe turns red after reopening the blocked vessels. 12/12 rats survived the procedure for more than one week. We are the first to report a surgical model for in vivo single liver lobe perfusion with a long survival period of more than one week. In contrast to the previously published report, the most important advantage of the technique presented here is that perfusion of 70% of the liver is maintained throughout the whole procedure. The establishment of this technique provides a foundation for in vivo partial liver engineering in rats, including decellularization and recellularization.

A Novel Surgical Technique As a Foundation for In Vivo Partial Liver Engineering in Rat

We establish a novel surgical technique for an in vivo single liver lobe perfusion model in rat as a prerequisite for further studying in vivo partial liver engineering in the future.