This work was supported by National Science Fund for Distinguished Young Scholars (81625002), National Natural Science Foundation of China (81470389, 81270282, 81601238), Program of Shanghai Academic Research Leader (18XD1402400), Shanghai Municipal Education Commission Gaofeng Clinical Medicine Grant Support (20152209), Shanghai Shenkang Hospital Development Center (16CR3034A), Shanghai Jiao Tong University (YG2013MS42), Shanghai Jiao Tong University School of Medicine (15ZH1003 and 14XJ10019), Shanghai Sailing Program (18YF1413000), and Postgraduate Innovation Program of Bengbu Medical College (Byycx1722). We thank Dr. Erhe Gao for his previous help in our lab.

All animal experiments were carried out according to the National Institutes of Health Guidelines on the Use of Laboratory Animals, and were approved by the Institute&#39;s Animal Ethics Committee. Male C57BL/6J mice (aged 8 - 10 weeks) were used for all the experiments. Mice were housed under pathogen-free conditions at 24 &#176;C &#177; 4 &#176;C, under a 12-h light/dark cycle, with free access to water and food.
1. Preparation of Adenovirus Solution
<ol>
	<li>Upon arrival of the purified adenovirus solution, store the solution in a -80 &#176;C freezer. 
	NOTE: The adenovirus injection was performed in viable mice. To ensure the sterility of the adenovirus vector, the adenovirus (encoding luciferase, VDR, or shRNA targeting VDR) were prepared and purified commercially9,10.</li>
	<li>On the day of operation, take out the purified adenovirus solution (3 x&#160;1010 plaque forming units [pfu]/mL) from the -80 &#176;C freezer, and thaw adenovirus vector solution on ice.</li>
	<li>Put on a facemask, sterile gloves, and sterile gown.</li>
	<li>Prepare a 50-&#181;L Hamilton syringe which has been sterilized the day before the operation. 
	NOTE: For sterilization, the Hamilton syringe was wrapped in gauze and placed in a high temperature/high pressure sterilizer. The mode of &#34;Sterilization for the solid&#34; was selected and sterilization was commenced by pressing the &#34;start&#34; button.</li>
	<li>After complete thawing of the purified adenovirus solution, spray the tube containing adenovirus solution with 75% ethanol, and move the adenovirus solution into a laminar flow sterile hood.</li>
	<li>In a laminar flow sterile hood, aspirate 50 &#181;L adenovirus solution using the Hamilton syringe without a needle, followed by attachment of a 30 G needle to the Hamilton syringe.</li>
	<li>Hold the syringe with the 30 G needle upward, and gently push the plunger of the syringe until the needle is filled with adenovirus solution (confirmed by the appearance of the first solution drops from the needle tip). 
	NOTE: It takes approximately 45 &#181;L solution to fill the 30 G needle, so now almost no solution is visible in the syringe.</li>
	<li>Use the above prepared syringe to carefully aspirate another 30 &#181;L adenovirus solution, and place the loosely capped syringe on ice for subsequent use.</li>
</ol>
2. Anesthesia and Operative Preparation
<ol>
	<li>Add 20 mL isoflurane into an isoflurane vaporizer, and connect the isoflurane vaporizer to an oxygen tank.</li>
	<li>Open the valve and let the oxygen flow from the oxygen tank to the isoflurane vaporizer. Maintain the oxygen flow rate at 2 L/min via an oxygen flow monitor in the isoflurane vaporizer.</li>
	<li>Induce anesthesia of the mouse with 4% isoflurane in 100% oxygen (2 L/min) for 2 min in a plastic cage connected to the isoflurane vaporizer.</li>
	<li>Take out the anesthetized mouse from the plastic cage, and secure it on a plastic platform in the prone position in a laminar flow sterile hood, and maintain anesthesia with 2% isoflurane in 100% oxygen (2 L/min) via a nose cone.</li>
	<li>Confirm adequate anesthesia by the absence of toe pinch response.</li>
	<li>Apply sterile ophthalmic cream to each eye to protect the corneas from drying.</li>
	<li>Shave the chest and the upper abdomen. Apply commercially available depilatory cream to the shaved site for 1 min. Remove the depilatory cream and remaining fur with wet gauzes.</li>
	<li>Sterilize the surgical site (left lower part of the chest) with 3 scrubs of iodine-chlorhexidine based antiseptic (e.g. entoiodine). Cover the surgical site with a sterile drape.</li>
</ol>
3. Intramyocardial Injection of Adenovirus in Mouse Heart
<ol>
	<li>Take off the gloves and put on a new pair of sterile gloves.</li>
	<li>Sterilize forceps, a micro-mosquito hemostat, a pair of surgical scissors, and a needle holder on the day before operation in a high temperature/high pressure sterilizer (see step 1.4 note).</li>
	<li>Make a 0.5-cm skin incision along the line connecting the xiphoid and axilla. Bluntly dissect the pectoral major and pectoral minor muscles with forceps and a micro-mosquito hemostat.</li>
	<li>Expose the intercostal spaces by retracting the pectoral major and pectoral minor muscles. Pierce through and open the fourth intercostal space (or the widest intercostal space by observation) with a micro-mosquito hemostat.</li>
	<li>Push the heart toward the incision to externalize the heart with the index finger of the non-dominant hand by gently pressing against the right side of the chest wall.</li>
	<li>Gently secure the externalized heart with the index finger and thumb of the non-dominant hand.</li>
	<li>Inject a total of 30 &#181;L adenovirus solution into the myocardium of the left ventricle in three sites (ventral, dorsal, and lateral wall of the left ventricle) via the Hamilton syringe filled with adenovirus (Figure 1C) with the dominant hand.</li>
	<li>After completion of injection, immediately place the heart back into the intrathoracic space.</li>
	<li>Manually evacuate the air in the intrathoracic space by gently pressing the chest wall toward the skin incision site. 
	Note: Successful air evacuation can be evidenced by flapping of the pectoral major and pectoral minor muscles caused by the expelled air.</li>
	<li>Close the skin by horizontal mattress suture with a 5-0 silk suture. 
	Note: The pectoral major and pectoral minor muscles should not be sutured, because they are only bluntly dissected and their anatomical structures are intact throughout the operation.</li>
</ol>
4. Postoperative Management
<ol>
	<li>Administer buprenorphine (3 mg/kg) twice daily via subcutaneous injection to reduce post-operative pain for the first 48 h after operation11.</li>
	<li>After operation, immediately maintain the mice on a heat pad (37 &#176;C) until fully recovered. Place the mouse back to the cage after it fully recovers.</li>
	<li>Sterilize the used Hamilton syringe and 30 G syringe needle in a high temperature/high pressure sterilizer (see step 1.4 note). Collect the sterilized 30 G syringe needle in a sharps container.</li>
</ol>
5. In Vivo Imaging for Measuring Cardiac Luciferase Expression
<ol>
	<li>On day 5 after adenovirus injection, prepare a fresh stock solution of luciferin at 15 mg/mL in Dulbecco&#39;s phosphate buffered saline (DPBS). Filter the solution through a 0.2 &#181;m filter.</li>
	<li>Intra-peritoneally inject the awake mice with the luciferin solution at 150 mg/kg body weight.</li>
	<li>Turn on the imaging system.</li>
	<li>After automatic self-test of the imaging system, select the imaging mode as &#34;Luminescent,&#34;&#160;and make the following settings: Exposure: Auto; Binning: 8; FStop: 1; Excitation: Block; Emission: Open.</li>
	<li>At 10 min after luciferin injection, anesthetize the mice by chloral hydrate injection (300 mg/kg body weight). Confirm adequate anesthesia by the absence of toe pinch response.</li>
	<li>Secure the anesthetized mice on a heated imaging stage in the prone position, with the chest directed towards the camera.</li>
	<li>Collect images (1 image for each mouse) and quantify the intensity of signals by drawing identical circular measurement regions of interest (ROI) around the chest.</li>
	<li>After the collection of images, sacrifice the mice and collect the tissues as mentioned in the following section.</li>
</ol>
6. Harvesting Tissues
<ol>
	<li>At different time points after virus injection (Figure 1B), anesthetize the mice with 4% isoflurane, and maintain anesthesia with 2% isoflurane via a nose cone, and secure it on a plastic platform in the prone position.</li>
	<li>Make a midline ventral incision in the anterior neck. Expose the right common carotid artery by retracting the omohyoid and sternocleidomastoid muscles.</li>
	<li>Sacrifice the mice by transecting the right common carotid artery and draining the blood.</li>
	<li>Make a midline ventral incision in the abdomen. Cut the ribs from both sides of the rib cage along the midclavicular line, and transect the diaphragm.</li>
	<li>Expose the heart by lifting the xiphoid. Find the ascending aorta and gently remove the perivascular fat tissue.</li>
	<li>Cannulate the aorta and retrogradely perfuse the heart with a 30 G needle attached to a 1 mL syringe filled with 0.5 mL of 4 &#176;C phosphate buffer solution (PBS).</li>
	<li>After perfusion, excise the heart, and divide it into the left and right ventricles. Excise the liver, lung, and spleen.</li>
	<li>After two washes in cold PBS (4 &#176;C), store all the collected tissues separately in cryogenic vials in liquid nitrogen.</li>
</ol>
7. Determination of Protein Expression
<ol>
	<li>Prepare homogenization buffer by adding protease inhibitor cocktail into commercially available lysis buffer at a ratio of 1:100. Calculate the total volume based on the number of samples to be processed. Keep the prepared homogenization buffer at 4 &#176;C for subsequent use.</li>
	<li>Take out the tissues from the liquid nitrogen tank. Weigh the tissue on high-precision scales (approximately 60 mg for each piece of tissue).</li>
	<li>Transfer the tissue to a new 1.5-mL microcentrifuge tube. Immediately add 200 &#181;L homogenization buffer and precooled 1.5-mm steel balls (4 &#176;C) into the microcentrifuge tube.</li>
	<li>Secure the microcentrifuge tubes in precooled (4 &#176;C) metal holders in an automatic tissue grinding machine, and commence homogenization by starting the machine with the following settings: Frequency: 70 Hz; time: 120 s.</li>
	<li>After homogenization, centrifuge the homogenate (16,000 &#215; g, 4 &#176;C), and carefully transfer the supernatant to a new 1.5-mL tube with a 100 &#181;L pipette.</li>
	<li>Add loading buffer solution (5x)&#160;to the protein supernatant at a ratio of 1:4, and denature the protein at 100 &#176;C for 5 min.</li>
	<li>The protein expression levels in the heart and other organ tissues are further monitored by Western blot analysis (representative results are shown in Figure 2C and Figure 3) according to the protocol previously described12.</li>
</ol>

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The authors have nothing to disclose.

The current report demonstrates a modified technique for intramyocardial injection of viral vectors for cardiac gene manipulation, which was modified from a method for myocardial infarction induction by Gao et al.13 Currently, in vivo characterization of specific gene functions most often involve the generation of knockout or transgenic mice3,14,15,16<s...

Cardiac disease is the leading cause of morbidity and mortality worldwide1,2. The lack of effective therapeutic strategies for life-threatening heart conditions including myocardial infarction and heart failure attracts intensive exploration of underlying pathomechanisms and identification of novel therapeutic options3. For these scientific explorations, cardiac-specific gene manipulation is widely used4,5. Cardiac gene manipulation can be achieved by genome editing using the powerful transcription activator-like effector nuclease (TALEN) and clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR associated protein 9 (Cas9) tools, or by delivery of ectopic genetic materials (e.g., virus vectors carrying genes encoding proteins of interest)6. Though genome editing allows precise and spatiotemporal genome modifications in living mice, it is still a time-consuming and labor-intensive practice6. Alternatively, cardiac-specific gene manipulation by virus vector or small interfering RNA (siRNA) complex delivery are routinely performed6.
Virus vector delivery to the adult mouse heart is achieved by roughly two strategies: systemic or local injection. Systemic injection of cardiotropic serotype of AAVs such as AAV9 is noninvasive for cardiac specific gene manipulation7. However, this method requires a relatively high amount of vector necessary for efficient transduction and gene expression, and may result in significant transduction of nontarget organs such as the muscle and liver7. Local virus injection is achieved by intramyocardial injection or intracoronary delivery7. Intracoronary delivery leads to a more even distribution of virus within the heart compared to intramyocardial injection. However, the disadvantages of this technique are the rapid wash out of viral vectors to the systemic circulation and transduction in nontarget organs8, and its requirement of devices for pressure measurement during the operation. By contrast, intramyocardial injection enables better virus retention in the myocardium as well as site specific delivery, but it fails to evenly distribute viral vector7. For small animals, intracoronary delivery is technically difficult to perform, while systemic AAV9 injection and intramyocardial injection are more commonly practiced4,5,7. Though systemic injection is easy to perform, conventional intramyocardial injection requires mechanical ventilation and thoracotomy, causes extensive tissue damage, and is time-consuming.
In this report, we described an easy, time-saving, and highly efficient method for intramyocardial injection. Adenovirus encoding luciferase and VDR, or shRNA targeting VDR, was injected to manipulate cardiac gene expression. Once mastered, this method can be used for gene manipulation, as well as injection of cells or other materials in the mouse heart.

<table><tbody><tr><td>&lt;strong&gt;Equipments&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Laminar flow sterile hood</td><td>Fengshi Animal Experimental&amp;nbsp; Equipment Techonology Co., Ltd. (Soochow, China)</td><td>FS-CJ-2F</td><td></td></tr><tr><td>Centrifuge</td><td>Thermo Scientific (Waltham, USA)</td><td>75005282</td><td></td></tr><tr><td>Tissue grinding machine</td><td>Scientz Biotechnology Co., Ltd. (Ningbo, China)</td><td>Scientz-48</td><td></td></tr><tr><td>High temperature/high pressure sterilizer</td><td>Hirayama (Saitama, Japan)</td><td>HVE-50</td><td></td></tr><tr><td>Isoflurane vaporizer&amp;nbsp;</td><td>Matrix (Orchard Park, USA)</td><td>VIP3000</td><td></td></tr><tr><td>IVIS&amp;nbsp; Lumina III imaging system</td><td>PerkinElmer (Waltham, USA)</td><td>CLS136334</td><td></td></tr><tr><td>Precision&amp;nbsp;balance</td><td>Sartorius (G&amp;ouml;ttingen, Germany)</td><td>28091873</td><td></td></tr><tr><td>&lt;strong&gt;Instruments&amp;nbsp;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Eppendorf pipette (100 &amp;micro;L)</td><td>Eppendorf (Westbury, USA)</td><td>&amp;nbsp;4920000059</td><td></td></tr><tr><td>Eppendorf pipette (10 &amp;micro;L)</td><td>Eppendorf (Westbury, USA)</td><td>&amp;nbsp;4920000113</td><td></td></tr><tr><td>Forceps</td><td>Shanghai Medical Instruments (Group) Ltd., Corp.&amp;nbsp;</td><td>JD4020</td><td>Curved tip</td></tr><tr><td>Hamilton syringe</td><td>Hamilton (Nevada, USA)</td><td>80501</td><td>Volume 50 &amp;mu;L</td></tr><tr><td>Micro-mosquito hemostat</td><td>F.S.T (Foster City, USA)</td><td>13011-12</td><td>Curved, tip width 1.3mm</td></tr><tr><td>Needle holder&amp;nbsp;</td><td>Shanghai Medical Instruments (Group) Ltd., Corp. (Shanghai, China)</td><td>J32110</td><td></td></tr><tr><td>Surgical scissors</td><td>F.S.T (Foster City, USA)</td><td>14002-12</td><td></td></tr><tr><td>1-mL Syringe</td><td>WeiGao Group Medical Polymer Co.,Ltd. (ShangDong, China)</td><td></td><td></td></tr><tr><td>&lt;strong&gt;Materials and reagents&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Anti-GAPDH antibody</td><td>CST (Danvers,&amp;nbsp; USA)</td><td>#2118</td><td></td></tr><tr><td>Anti-Luciferase antibody</td><td>Abcam (Cambridge, UK)</td><td>ab187340</td><td></td></tr><tr><td>Anti-rabbit IgG</td><td>CST (Danvers,&amp;nbsp; USA)</td><td>&amp;nbsp;#7074</td><td></td></tr><tr><td>Anti-VDR antibody</td><td>Abcam (Cambridge, UK)</td><td>&amp;nbsp;ab109234</td><td></td></tr><tr><td>Buprenorphine</td><td>Thermo Scientific (Waltham, USA)</td><td>PA175056</td><td></td></tr><tr><td>Chloralic hydras</td><td>LingFeng Chemical (ShangHai, China)</td><td></td><td></td></tr><tr><td>Cryogenic Vials</td><td>Thermo Scientific (Waltham, USA)</td><td>375418</td><td>1.8 mL&amp;nbsp;</td></tr><tr><td>Depilatory cream</td><td>Veet (Shanghai, China)</td><td></td><td></td></tr><tr><td>Dulbecco&#039;s phosphate buffered saline&amp;nbsp;</td><td>Gibco (Grand Island,&amp;nbsp; USA)</td><td>14040133</td><td></td></tr><tr><td>Entoiodine</td><td>LiKang (Shanghai, China)</td><td>310132</td><td></td></tr><tr><td>EP tube</td><td>Sarstedt (Newton, USA)</td><td>PCR001</td><td></td></tr><tr><td>Filter</td><td>Millipore (Bedford, USA)</td><td></td><td>Pore size 0.2 &amp;micro;m&amp;nbsp;</td></tr><tr><td>Isoflurane</td><td>Yipin Pharmaceutical Company (Hebei, China)</td><td></td><td></td></tr><tr><td>Luciferin</td><td>Promega (Madison, USA)</td><td>P1041</td><td></td></tr><tr><td>Lysis buffer for western&amp;nbsp; blot</td><td>Beyotime (Shanghai, China)</td><td>P0013J</td><td>Without inhibitors</td></tr><tr><td>Ophthalmic cream</td><td>Apex Laboratories ( Melbourne, Australia))</td><td></td><td></td></tr><tr><td>PBS</td><td>Gibco (Grand Island,&amp;nbsp; USA)</td><td>10010023</td><td></td></tr><tr><td>Protease inhibitor cocktail</td><td>Thermo Scientific (Waltham, USA)</td><td>78438</td><td></td></tr><tr><td>5-0 silk suture</td><td>Shanghai Medical Instruments (Group) Ltd., Corp. (Shanghai, China)</td><td></td><td></td></tr><tr><td>Steel ball</td><td>Scientz Biotechnology Co., Ltd. (Ningbo, China)</td><td></td><td>Width 1.5 mm</td></tr><tr><td>Syringe needle</td><td>Kindly Medical Devices Co., Ltd. (Zhejiang, China)</td><td></td><td>30 gauge&amp;nbsp;</td></tr><tr><td>Warm mat</td><td>Warmtact Electrical Heating Technology Co., Ltd. (Guangdong, China )</td><td>NF-GNCW</td><td></td></tr></tbody></table>

a simple and efficient method for in vivo cardiac-specific gene manipulation by intramyocardial injection in mice

Gene manipulation specifically in the heart significantly potentiate the investigation of cardiac disease pathomechanisms and their therapeutic potential. In vivo cardiac-specific gene delivery is commonly achieved by either systemic or local delivery. Systemic injection via tail vein is easy and efficient in manipulating cardiac gene expression by using recombinant adeno-associated virus 9 (AAV9). However, this method requires a relatively high amount of vector for efficient transduction, and may result in nontarget organ gene transduction. Here, we describe a simple, efficient, and time-saving method of intramyocardial injection for in vivo cardiac-specific gene manipulation in mice. Under anesthesia (without ventilation), the pectoral major and minor muscles were bluntly dissected, and the mouse heart was quickly exposed by manual externalization through a small incision at the fourth intercostal space. Subsequently, adenovirus encoding luciferase (Luc) and vitamin D receptor (VDR), or short hairpin RNA (shRNA) targeting VDR, was injected with a Hamilton syringe into the myocardium. Subsequent in vivo imaging demonstrated that luciferase was successfully overexpressed specifically in the heart. Moreover, Western blot analysis confirmed the successful overexpression or silencing of VDR in the mouse heart. Once mastered, this technique can be used for gene manipulation, as well as injection of cells or other materials such as nanogels in the mouse heart.

The experiment protocol and some of the key steps for the reported method are shown in Figure 1. At 5 days after intramyocardial injection of adenovirus encoding luciferase (Adv-luc), in vivo imaging in adv-luc injected mice indicated robust overexpression of luciferase specifically in the heart (Figure 2A, B), which was confirmed by Western blot analysis (Figure 2C), sugges...

Watch this Scientific Journal Video about A Simple and Efficient Method for In Vivo Cardiac-specific Gene Manipulation by Intramyocardial Injection in Mice at JoVE.com

A Simple and Efficient Method for In Vivo Cardiac-specific Gene Manipulation by Intramyocardial Injection in Mice

Here we present a protocol for cardiac-specific gene manipulation in mice. Under anesthesia, the mouse hearts were externalized through the fourth intercostal space. Subsequently, adenoviruses encoding specific genes were injected with a syringe into the myocardium, followed by protein expression measurement via in vivo imaging and Western blot analysis.

A Simple and Efficient Method for In Vivo Cardiac-specific Gene Manipulation by Intramyocardial Injection in Mice

Gene manipulation specifically in the heart significantly potentiate the investigation of cardiac disease pathomechanisms and their therapeutic potential. ...

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Research

JoVE Journal

Biology

15.9K Views.  The First Affiliated Hospital of Bengbu Medical College. Here we present a protocol for cardiac-specific gene manipulation in mice. Under anesthesia, the mouse hearts were externalized through the fourth intercostal space. Subsequently, adenoviruses encoding specific genes were injected with a syringe into the myocardium, followed by protein expression measurement via in vivo imaging and Western blot analysis.

Video: A Simple and Efficient Method for In Vivo Cardiac-specific Gene Manipulation by Intramyocardial Injection in Mice

Coronary Artery Ligation and Intramyocardial Injection in a Murine Model of Infarction

Mouse models are a valuable tool for studying acute injury and chronic remodeling of the myocardium in vivo. With the advent of genetic modifications to the whole organism or the myocardium and an array of biological and/or synthetic materials, there is great potential for any combination of these to assuage the extent of acute ischemic injury and impede the onset of heart failure pursuant to myocardial remodeling. 
Here we present the methods and materials used to reliably perform this microsurgery and the modifications involved for temporary (with reperfusion) or permanent coronary artery occlusion studies as well as intramyocardial injections. The effects on the heart that can be seen during the procedure and at the termination of the experiment in addition to histological evaluation will verify efficacy. 
Briefly, surgical preparation involves anesthetizing the mice, removing the fur on the chest, and then disinfecting the surgical area. Intratracheal intubation is achieved by transesophageal illumination using a fiber optic light. The tubing is then connected to a ventilator. An incision made on the chest exposes the pectoral muscles which will be cut to view the ribs. For ischemia/reperfusion studies, a 1 cm piece of PE tubing placed over the heart is used to tie the ligature to so that occlusion/reperfusion can be customized. For intramyocardial injections, a Hamilton syringe with sterile 30gauge beveled needle is used. When the myocardial manipulations are complete, the rib cage, the pectoral muscles, and the skin are closed sequentially. Line block analgesia is effected by 0.25% marcaine in sterile saline which is applied to muscle layer prior to closure of the skin. The mice are given a subcutaneous injection of saline and placed in a warming chamber until they are sternally recumbent. They are then returned to the vivarium and housed under standard conditions until the time of tissue collection. At the time of sacrifice, the mice are anesthetized, the heart is arrested in diastole with KCl or BDM, rinsed with saline, and immersed in fixative. Subsequently, routine procedures for processing, embedding, sectioning, and histological staining are performed. 
Nonsurgical intubation of a mouse and the microsurgical manipulations described make this a technically challenging model to learn and achieve reproducibility. These procedures, combined with the difficulty in performing consistent manipulations of the ligature for timed occlusion(s) and reperfusion or intramyocardial injections, can also affect the survival rate so optimization and consistency are critical.

Numerous genetic manipulations and/or intramyocardial injections of genes, proteins, cells, and/or biomaterials are superimposed upon the dimension of time in studies of acute ischemia/ reperfusion injury and chronic remodeling in mice. This video illustrates the microsurgical procedures for ischemia/reperfusion, permanent coronary artery ligation, and intramyocardial injection studies.

Mouse models are a valuable tool for studying acute injury and chronic remodeling of the myocardium in vivo.  With the advent of genetic modifications to ...

Medicine

Delivery of Modified mRNA in a Myocardial Infarction Mouse Model

Myocardial infarction (MI) is a leading cause of morbidity and mortality in the Western world. In the past decade, gene therapy has become a promising treatment option for heart disease, owing to its efficiency and exceptional therapeutic effects. In an effort to repair the damaged tissue post-MI, various studies have employed DNA-based or viral gene therapy but have faced considerable hurdles due to the poor and uncontrolled expression of the delivered genes, edema, arrhythmia, and cardiac hypertrophy. Synthetic modified mRNA (modRNA) presents a novel gene therapy approach that offers high, transient, safe, nonimmunogenic, and controlled mRNA delivery to the heart tissue without any risk of genomic integration. Due to these remarkable characteristics combined with its bell-shaped pharmacokinetics in the heart, modRNA has become an attractive approach for the treatment of heart disease. However, to increase its effectiveness in vivo, a consistent and reliable delivery method needs to be followed. Hence, to maximize modRNA delivery efficiency and yield consistency in modRNA use for in vivo applications, an optimized method of preparation and delivery of modRNA intracardiac injection in a mouse MI model is presented. This protocol will make modRNA delivery more accessible for basic and translational research.

This protocol presents a simple and coherent way to transiently upregulate a gene of interest using modRNA after myocardial infarction in mice.

Myocardial infarction (MI) is a leading cause of morbidity and mortality in the Western world. In the past decade, gene therapy has become a promising ...

Genetics

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 ...

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a general challenge in developmental biology research. In the embryonic heart, this can be particularly difficult as regional differences in the expression of transcriptional regulators, paracrine/juxtacrine signaling cues, and hemodynamic force are all known to influence cardiomyocyte maturation. A simplified method to alter a developing cardiomyocyte's molecular and biomechanical microenvironment would, therefore, serve as a powerful technique to examine how local conditions influence cell fate and function. To address this, we have optimized a method to physically transplant juvenile cardiomyocytes into ectopic locations in the heart or the surrounding embryonic tissue. This allows us to examine how microenvironmental conditions influence cardiomyocyte fate transitions at single cell resolution within the intact embryo. Here, we describe a protocol in which embryonic myocytes can be isolated from a variety of cardiac sub-domains, dissociated, fluorescently labeled, and microinjected into host embryos with high precision. Cells can then be directly analyzed in situ using a variety of imaging and histological techniques. This protocol is a powerful alternative to traditional grafting experiments that can be prohibitively difficult in a moving tissue such as the heart. The general outline of this method can also be adapted to a variety of donor tissues and host environments, and its ease of use, low cost, and speed make it a potentially useful application for a variety of developmental studies.

In this method, embryonic cardiac tissues are surgically microdissected, dissociated, fluorescently labeled, and implanted into host embryonic tissues. This provides a platform for studying the individual or tissue level developmental organization under ectopic hemodynamic conditions, and/or altered paracrine/juxtacrine environments.

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a ...

Developmental Biology

Ultrasound-guided Transthoracic Intramyocardial Injection in Mice

Murine models of cardiovascular disease are important for investigating pathophysiological mechanisms and exploring potential regenerative therapies. Experiments involving myocardial injection are currently performed by direct surgical access through a thoracotomy. While convenient when performed at the time of another experimental manipulation such as coronary artery ligation, the need for an invasive procedure for intramyocardial delivery limits potential experimental designs. With ever improving ultrasound resolution and advanced noninvasive imaging modalities, it is now feasible to routinely perform ultrasound-guided, percutaneous intramyocardial injection. This modality efficiently and reliably delivers agents to a targeted region of myocardium. Advantages of this technique include the avoidance of surgical morbidity, the facility to target regions of myocardium selectively under ultrasound guidance, and the opportunity to deliver injectate to the myocardium at multiple, predetermined time intervals. With practiced technique, complications from intramyocardial injection are rare, and mice quickly return to normal activity on recovery from anesthetic. Following the steps outlined in this protocol, the operator with basic echocardiography experience can quickly become competent in this versatile, minimally invasive technique.

Echocardiography-guided percutaneous intramyocardial injection represents an efficient, reliable, and targetable modality for the delivery of gene transfer agents or cells into the murine heart. Following the steps outlined in this protocol, the operator can quickly become competent in this versatile, minimally invasive technique.

Murine models of cardiovascular disease are important for investigating pathophysiological mechanisms and exploring potential regenerative therapies. ...

Electroporation-Based Transgenesis of Cardiac Cells in Mouse Hearts

An Efficient Transgenesis Approach for Gene Delivery in the Mouse Embryonic Heart

The mammalian heart is a complex organ formed during development via highly diverse populations of progenitor cells. The origin, timing of recruitment, and fate of these progenitors are vital for the proper development of this organ. The molecular mechanisms that govern the morphogenesis of the heart are essential for understanding the pathogenesis of congenital heart diseases and embryonic cardiac regeneration. Classical approaches to investigate these mechanisms employed the generation of transgenic mice to assess the function of specific genes during cardiac development. However, mouse transgenesis is a complex, time-consuming process that often cannot be performed to assess the role of specific genes during heart development. To address this, we have developed a protocol for efficient electroporation and culture of mouse embryonic hearts, enabling transient transgenesis to rapidly assess the effect of gain- or loss-of-function of genes involved in cardiac development. Using this methodology, we successfully overexpressed Meis1 in the embryonic heart, with a preference for epicardial cell transfection, demonstrating the capabilities of the technique.

Author Spotlight: Cardiac Cell Transgenesis for Rapid Gene Screening

This protocol presents a detailed methodological framework for electroporation-based transgenesis of cardiac cells in developing mouse hearts. The video assets provided here will facilitate learning of this versatile technique.

The mammalian heart is a complex organ formed during development via highly diverse populations of progenitor cells. The origin, timing of recruitment, ...

Percutaneous Contrast Echocardiography-guided Intramyocardial Injection and Cell Delivery in a Large Preclinical Model

Cell and gene therapy are exciting and promising strategies for the purpose of cardiac regeneration in the setting of heart failure with reduced ejection fraction (HFrEF). Before they can be considered for use, and implemented in humans, extensive preclinical studies are required in large animal models to evaluate the safety, efficacy, and fate of the injectate (e.g., stem cells) once delivered into the myocardium. Small rodent models offer advantages (e.g., cost effectiveness, amenability for genetic manipulation); however, given inherent limitations of these models, the findings in these rarely translate into the clinic. Conversely, large animal models such as rabbits, have advantages (e.g., similar cardiac electrophysiology compared to humans and other large animals), whilst retaining a good cost-effective balance. Here, we demonstrate how to perform a percutaneous contrast echocardiography-guided intramyocardial injection (IMI) technique, which is minimally invasive, safe, well tolerated, and very effective in the targeted delivery of injectates, including cells, into several locations within the myocardium of a rabbit model. For the implementation of this technique, we also have taken advantage of a widely available clinical echocardiography system. After putting in practice the protocol described here, a researcher with basic ultrasound knowledge will become competent in the performance of this versatile and minimally invasive technique for routine use in experiments, aimed at hypothesis testing of the capabilities of cardiac regenerative therapeutics in the rabbit model. Once competency is achieved, the whole procedure can be performed within 25 min after anaesthetizing the rabbit.

Novel therapeutic strategies in cardiac regenerative medicine require extensive and detailed studies in large preclinical animal models before they can be considered for use in humans. Here, we demonstrate a percutaneous contrast echocardiography-guided intramyocardial injection technique in rabbits, which is valuable for hypothesis testing the efficacy of such novel therapies.

Cell and gene therapy are exciting and promising strategies for the purpose of cardiac regeneration in the setting of heart failure with reduced ejection ...

Adeno-Associated Virus-Mediated Delivery of CRISPR for Cardiac Gene Editing in Mice

The clustered, regularly interspaced, short, palindromic repeat (CRISPR) system has greatly facilitated genome engineering in both cultured cells and living organisms from a wide variety of species. The CRISPR technology has also been explored as novel therapeutics for a number of human diseases. Proof-of-concept data are highly encouraging as exemplified by recent studies that demonstrate the feasibility and efficacy of gene editing-based therapeutic approach for Duchenne muscular dystrophy (DMD) using a murine model. In particular, intravenous and intraperitoneal injection of the recombinant adeno-associated virus (rAAV) serotype rh.74 (rAAVrh.74) has enabled efficient cardiac delivery of the Staphylococcus aureus CRISPR-associated protein 9 (SaCas9) and two guide RNAs (gRNA) to delete a genomic region with a mutant codon in exon 23 of mouse Dmd gene. This same approach can also be used to knock out the gene-of-interest and study their cardiac function in postnatal mice when the gRNA is designed to target the coding region of the gene. In this protocol, we show in detail how to engineer rAAVrh.74-CRISPR vector and how to achieve highly efficient cardiac delivery in neonatal mice.

Here we provide a detailed protocol to carry out in vivo cardiac gene editing in mice using recombinant Adeno-Associated Virus(rAAV)-mediated delivery of CRISPR. This protocol offers a promising therapeutic strategy to treat dystrophic cardiomyopathy in Duchenne muscular dystrophy and can be used to generate cardiac-specific knockout in postnatal mice.

The clustered, regularly interspaced, short, palindromic repeat (CRISPR) system has greatly facilitated genome engineering in both cultured cells and ...

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect their in vivo potential. Preferably, these cells should be placed in a proper embryonic environment in order to acquire their definite phenotype. Furthermore, cell lineage tracing studies in the mouse after labeling cells with dyes or retroviral vectors has remained mostly limited to early stage mouse embryos with still poorly developed organs. To overcome these limitations, we designed standard and ultrasound-mediated microinjection protocols to inject various agents in targeted regions of the heart in mouse embryos at E9.5 and later stages of development. &#160;Embryonic explant or embryos are then cultured or left to further develop in utero. These agents include fluorescent dyes, virus, shRNAs, or stem cell-derived progenitor cells. Our approaches allow for preservation of the function of the organ while monitoring migration and fate of labeled and/or injected cells. These technologies can be extended to other organs and will be very helpful to address key biological questions in biology of development.

We describe a series of methods to inject dyes, DNA vectors, virus, and cells in order to monitor both cell fate and phenotype of endogenous and grafted cells derived from embryonic or pluripotent cells within mouse embryos at embryonic day (E)9.5 and later stages of development.

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect ...

Ultrasound-Guided Induced Pluripotent Stem Cell-Derived Cardiomyocyte Implantation in Myocardial Infarcted Mice

The key objective of cell therapy after myocardial infarction (MI) is to effectively enhance the cell grafted rate, and human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are a promising cell source for cardiac repair after ischemic damage. However, a low grafted rate is a significant obstacle for effective cardiac tissue regeneration after transplantation. This protocol shows that multiple hiPSC-CM ultrasound-guided percutaneous injections into an MI area effectively increase cell transplantation rates. The study also describes the entire hiPSC-CM culture process, pretreatment, and ultrasound-guided percutaneous delivery methods. In addition, the use of human mitochondrial DNA help detect the absence of hiPSC-CMs in other mouse organs. Lastly, this paper describes the changes in cardiac function, angiogenesis, cell size, and apoptosis at the infarcted border zone in mice 4 weeks after cell delivery. It can be concluded that echocardiography-guided percutaneous injection of the left ventricular myocardium is a feasible, relatively invasive, satisfactory, repeatable, and effective cellular therapy.

Ultrasound-guided cell delivery around the site of myocardial infarction in mice is a safe, effective, and convenient way of cell transplantation.

The key objective of cell therapy after myocardial infarction (MI) is to effectively enhance the cell grafted rate, and human induced pluripotent stem ...

A Simple and Efficient Method for In Vivo Cardiac-specific Gene Manipulation by Intramyocardial Injection in Mice

Podsumowanie

Przeglądaj więcej filmów

Coronary Artery Ligation and Intramyocardial Injection in a Murine Model of Infarction

Delivery of Modified mRNA in a Myocardial Infarction Mouse Model

Intramyocardial Cell Delivery: Observations in Murine Hearts

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Ultrasound-guided Transthoracic Intramyocardial Injection in Mice

An Efficient Transgenesis Approach for Gene Delivery in the Mouse Embryonic Heart

Percutaneous Contrast Echocardiography-guided Intramyocardial Injection and Cell Delivery in a Large Preclinical Model

Adeno-Associated Virus-Mediated Delivery of CRISPR for Cardiac Gene Editing in Mice

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

Ultrasound-Guided Induced Pluripotent Stem Cell-Derived Cardiomyocyte Implantation in Myocardial Infarcted Mice