This study was supported by a grant from JSPS KAKENHI (Grant No. 23390072 and 19K07335) and AMED (Grant No. A-149).

The animal experiments were conducted according to the Kansai-Medical University ethical guidelines for animal experiments and were approved by the ethical committees (approval number: 23-104). All the animals were raised under a constant light-dark cycle in a specific pathogen-free environment. All the sterilized surgical tools, such as scissors, forceps, and retractors, were autoclaved and dried thoroughly.
1. Preparation of the neonatal rat cardiomyocyte balls
<ol>
	<li>Neonatal rat heart collection
	<ol>
		<li>For the heart collection, follow a procedure similar to that described in a previous report7.</li>
		<li>Immerse neonatal (0-2 days after birth) Sprague-Dawley (SD) rats in povidone-iodine and 70% ethanol solutions sequentially, and then transfer them into an air-tight case filled with vaporized isoflurane (the concentration should be over 10% v/v) for deep anesthesia.</li>
		<li>After the confirmation of unconsciousness by a loss of locomotor activity, decapitate the ratwhile holding the body from the back with the hand, and then cut 2-4 mm of tissue from the frontal center of the ribcage to the caudal and then in the rostral direction using sharp scissors.</li>
		<li>Grip the back skin to pull the cut open and push out the heart from the ribcage. Cut the ventricles using scissors and immerse them in phosphate-buffered saline (PBS) without calcium or magnesium (PBS(&#8722;)).</li>
	</ol>
	</li>
	<li>Dispersion of the neonatal rat cardiomyocytes
	<ol>
		<li>Mince the collected ventricles dispersed in a minimal amount of Ads buffer (Ads buffer: 116 mM NaCl, 20 mM HEPES, 12.5 mM NaH2PO4, 5.6 mM glucose, 5.4 mM KCl, 0.8 mM MgSO4, pH 7.35) in an autoclaved concave glassware into small pieces (1 mm x 1 mm) using curved scissors.</li>
		<li>Transfer the minced tissues and a micro-magnetic stirrer into a 50 mL centrifugation tube, and disperse the tissues into single cells with 0.1% collagenase, 0.1% trypsin, 20 &#181;g/mL DNase I, and 50 nM tetramethylrhodamine methyl ester in Ads buffer at 37 &#176;C by stirring for 30 min.</li>
		<li>Separate the aggregates and dispersed cells through natural sedimentation, collect only the dispersed cells into a tube, and digest the residual cell aggregates again with the same digestion medium.</li>
		<li>Continue this procedure until all the cells are completely dissociated. To confirm the complete dispersion of the cells, observe the tubes under a microscope (4x objective lens).</li>
		<li>Collect the dispersed cells using centrifugation at 150 x g for 5 min and dissociate them in 1-2 mL of Ads buffer.</li>
	</ol>
	</li>
	<li>Fluorescence-activated cell sorting of cardiomyocytes
	<ol>
		<li>Analyze the cells using fluorescence-activated cell sorting (FACS) using 556-601 nm bandpass filters to detect red fluorescence signals.</li>
		<li>Carefully perform pre-gating to eliminate doublet fractions14. The gate settings for doublet elimination should be as per the manufacturer&#39;s instructions.</li>
		<li>Set the forward scatter on the x-axis and the red fluorescence signal on the y-axis. Three populations were observed: a lowermost population containing erythrocytes and dead cells, a middle population containing non-cardiomyocytes, including fibroblasts and endothelial cells, and a top population containing pure ventricular cardiomyocytes7.</li>
	</ol>
	</li>
	<li>Preparation of red fluorescence-labeled cardiomyocyte balls
	<ol>
		<li>Selectively sort the cardiomyocytes, and centrifuge for 5 min at 150 x g. Completely dissolve the cell pellets in 1 mL of alpha-modified minimal essential medium (alpha-MEM) containing 10% heat-inactivated fetal bovine serum (FBS).</li>
		<li>Measure the cell concentration using a hemocytometer and dilute the cell suspension solution to 3,000 cells/mL with alpha-MEM 10% FBS.</li>
		<li>Distribute them into cell non-adhesive 96-well plates (100 &#181;L per well), centrifuge for 5 min at 100 x g, and culture for 2-3 d in a cell culture incubator with 5% CO2 at 37 &#176;C.</li>
		<li>Before the injection experiments, harvest a cardiomyocyte ball from each well via aspiration with the culture medium using a 1,000 &#181;L pipette, and collect them in a 15 mL tube.</li>
		<li>Stain with PKH26 following the manufacturer&#39;s instructions for tracking after engraftment.</li>
	</ol>
	</li>
</ol>
2. Preparations for the slow injection method
<ol>
	<li>Preparation of the gelatin stock solution
	<ol>
		<li>Weigh the gelatin, dissolve it in Ads buffer to produce a 10% w/v solution, and autoclave it.</li>
	</ol>
	</li>
	<li>Production of a cell injection device
	<ol>
		<li>Overall device design: Prepare the apparatus shown in Figure 1. This system is combined with a syringe cooling apparatus and main injection apparatus.</li>
		<li>Prepare the main injection apparatus described below:
		<ol>
			<li>For a neonatal cardiomyocyte ball injection, use a 29 G, 50 mm long needle equipped with a syringe.</li>
			<li>Connect a power transfer syringe (18G, 1 mL) back-to-back using a cell injection syringe (Figure 1). Connect the needle of the power transfer syringe to a thin polypropylene tube with low lumen expandability.</li>
			<li>Then, connect the other side of the power transfer tube to the needle of the same syringe (18G, 1 mL), and set it in a syringe pump. Fill the two power transfer syringes and tubes with water without any air bubbles. 
			NOTE: When one plunger of the power transfer syringe is pushed in, the pressure is directly transferred to the other syringe, and the plunger protrudes.</li>
		</ol>
		</li>
		<li>Set up the injection syringe cooling system as described below.
		<ol>
			<li>Wind a copper tube (external diameter = 1 mm; internal diameter = 0.3 mm; thickness = 0.35 mm) tightly around the cell-containing part of the cell injection syringe, leaving 10 mm of excess pipe at both ends.</li>
			<li>Connect the copper tube to flexible plastic tubing. Further, connect the other ends of the plastic tubing to an external pump, fill the line with cooling water, and cool the water by immersing the excess copper tube into crushed ice (Figure 1). 
			NOTE: This cooling system maintains the cell suspension solution in the cylinder at approximately 2 &#176;C.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
3. Development of a rat myocardial infarction model by obtuse coronary artery occlusion
<ol>
	<li>Anesthetize male immune-deficient nude rats (F344/Njcl-rnu/rnu) with air containing 3% isoflurane. Insert a cannula into the trachea and connect it to the respirator.</li>
	<li>Connect a cannula to an isoflurane vaporizer with a controller to maintain an 3% isoflurane concentration and, thus, sustain sufficient anesthesia. Confirm no response to pain stimuli. Apply veterinarian ointment on the eyes to prevent dryness.</li>
	<li>Fix the limbs with cut surgical tapes on a 40 &#176;C warmed surgical plate. Twist the body of the rat to the right of the body axis and use the left armpit as the surgical field.</li>
	<li>Remove the hair in the surgical field using a depilatory cream and wipe the skin with povidone-iodine. Using sharp scissors, cut a 1.5 cm incision in the skin and pectoralis major muscle.</li>
	<li>Confirm the third intercostal space and rip the intercostal muscles and costal pleura using micro forceps with blunt tips. Keep the chest open by using a retractor. Gently remove the thin pericardium with forceps.</li>
	<li>To construct an infarction model of the lateral wall of the heart, find the position 1 mm caudal to the tip of the left atrium to identify the obtuse coronary artery, pass a 7-0 silk suture, scoop up tissue that is 2.5 mm wide and 2.5 mm deep from dorsal to ventral, and ligate the tissue tightly.</li>
	<li>Confirm successful ligation by the weak contraction distal from the ligature. After gently removing the retractor, place a 5-0 silk suture between the second and fourth intercostal spaces, and close the thoracotomy.</li>
	<li>Decrease the isoflurane concentration to 1%. Gently suture the muscle and skin with 5-0 silk. Decrease the isoflurane concentration to 0% and wait for approximately 5 min until spontaneous breathing starts.</li>
	<li>Topically apply 2 mg/mL lidocaine in saline to the incision. Administrate 1 mL of saline via a subcutaneous injection. Apply veterinarian ointment topically to prevent infections.</li>
	<li>Remove the rat from the intubation tube, and return to the animal cage; then, raise the rats in individual cages for 1 week.</li>
	<li>Analyze the changes in the cardiac systolic pump function using echocardiography. 
	&#8203;NOTE: The cardiac function will be reduced owing to lateral myocardial infarction.</li>
</ol>
4. Echo-guided percutaneous cell transplantation using the slow injection method
<ol>
	<li>Pre-warm 10% gelatin stock at 37 &#176;C until it becomes liquid.</li>
	<li>Dilute 10% gelatin stock with pre-warmed Ads buffer to obtain the final injectable 5% w/v gelatin solution (100 &#181;L is required per animal).</li>
	<li>Suspend 96 cardiomyocyte balls (total: 28,800 cardiomyocytes/animal) in 100 &#181;L of pre-warmed injection solution.</li>
	<li>Load the suspension prepared in step 4.3 into the cell injection syringe, avoiding the aspiration of excess air.</li>
	<li>To eliminate bubbles in the syringe, hold it vertically with the needle upward, tap the syringe, and collect any bubbles at the upper ridge of the syringe. 
	NOTE: During this step, observe the cardiomyocyte balls gradually settle onto the rubber seal of the plunger.</li>
	<li>Maintain the vertical position of the syringe, push up the plunger slowly, and discard the bubbles and excess cell suspension solution until 20 &#181;L of the cell suspension remains in the syringe. Push the plunger carefully at a constant speed so that the cardiomyocyte ball remains resting on the rubber seal of the plunger.</li>
	<li>Immerse the capped syringe directly in an ice bath for 5 min.</li>
	<li>Place a cooled syringe in the injection apparatus. Tightly fix the injection syringe apparatus settled on a fine movement device on the animal stage using an X-Y-Z position-adjustable handmade clamp. The fine movement device can move the needle position using the x-direction 20 mm slide, y-direction swing, and z-direction bowing movements.</li>
	<li>Anesthetize the rats in a sealed box filled with air containing 3% isoflurane. Confirm anesthesia by no response to pain stimuli. Apply veterinarian ointment on the eyes to prevent dryness.</li>
	<li>Fix the limbs with cut surgical tapes on a 40 &#176;C warmed echo plate. To sustain sufficient anesthesia, ensure the inhaled air contains an approximately 3% concentration of isoflurane.</li>
	<li>Remove the hair at the injection field (2 cm in diameter) and chest with depilatory cream and wipe the skin with povidone-iodine.</li>
	<li>Apply echo-gel on the chest and echo probe. Place the echo probe close to the chest along the cranial and caudal axes and start to obtain B-mode echocardiography following the manufacturer&#39;s manual.</li>
	<li>Advance the tip of the injection needle into the myocardium at the frontal view (Figure 2 and Supplementary Video 1).</li>
	<li>To activate the main syringe pump, push the Start button, and rotate the dial to adjust to a predetermined number for an injection speed of approximately 0.02 &#181;L/s. Apply veterinarian ointment topically around the needling position to prevent infections. 
	NOTE: It is necessary to determine the appropriate dial number for the intended injection speed using an injection solution. After the injection, remove the injection needle using an exemplary movement system.</li>
	<li>Decrease the isoflurane concentration to 0% and wait for approximately 5 min until the animal regains sufficient consciousness to maintain sternal recumbency.</li>
</ol>
5. Evaluation of cardiac function
<ol>
	<li>Anesthetize the rats in a sealed box filled with air containing 3% isoflurane. Confirm anesthesia by no response to pain stimuli. Apply veterinarian ointment on the eyes to prevent dryness.</li>
	<li>Apply electrifying cream for ECG acquisition on the limb tips. Fix the limbs with cut surgical tapes on a 40 &#176;C warmed echo plate. Use a physiological monitoring system to detect real-time ECG and heart rate.</li>
	<li>According to the manufacturer&#39;s instructions, first determine the long-axis angle using the B-mode echo image showing the left ventricular apex to the outflow tract, and then rotate the echo-probe 90&#176; to change to the short-axis view.</li>
	<li>Using only the caudal to rostral axis fine movement system of the animal stage, adjust the short-axis view to the papillary muscle level. Then, change the image mode to M-mode by pressing the M-mode button, and record a video for 5 s by pressing the Cine-loop button.</li>
	<li>To analyze and calculate the fraction shortening using the software, press the Measure button and a vertical line tool to define the end-systolic and end-diastolic left ventricular internal dimensions. The software automatically calculates the percent fraction shortening (FS).</li>
	<li>Decrease the isoflurane concentration to 0% and wait for approximately 5 min until the animal regains sufficient consciousness to maintain sternal recumbency.</li>
</ol>
6. Immunohistochemistry
<ol>
	<li>Fix the euthanized body (perform euthanasia as in step 1.1.3) on the table using the cut surgical tapes, excise the hearts, wash with PBS, and immerse the heart in 4% paraformaldehyde/PBS.</li>
	<li>Dissect the hearts into three sections, immerse them in 40% sucrose for cryoprotection, embed in an optimal cutting temperature (OCT) compound, and freeze them at &#8722;80 &#176;C.</li>
	<li>Attach the cryosectioned tissues (8 &#181;m thickness) to aminosilane-coated glass slides. After sufficient drying using non-warmed wind generated by a general hair dryer, immerse the glass slides in tris-buffered saline containing 0.2% Tween-20 (TBS-T).</li>
	<li>Immerse them in a blocking solution for 30 min at 25 &#176;C. Pour 100 &#181;L of the primary antibody-containing blocking solution onto the slide, and incubate overnight at 4 &#176;C with paraffin sealing to keep the antibody solution spread on the tissue. 
	NOTE: The antibody concentration is shown in the Table of Materials.</li>
	<li>Lay the slides horizontally in a handmade high-humidity box utilizing the natural vapor from wet paper to prevent evaporation. Wash the slides three times with TBS-T.</li>
	<li>Treat with the secondary antibody-containing blocking agent for 1 h at 25 &#176;C in the same manner as the primary antibody. After three washes, observe the fluorescence signals using a fluorescence microscope and confocal laser microscope. 
	NOTE: The antibody concentration is shown in the Table of Materials.</li>
	<li>Statistical analysis: For the cell volume comparison, as shown in Figure 3A, perform a non-paired t-test; to assess the cardiac functional recovery following cardiomyocyte administration using the slow injection method, as shown in Figure 4A, use a paired t-test. In this work, differences were considered statistically significant at P &#60; .01. Error bars represent the standard deviation.</li>
</ol>

A Novel Minimally Invasive Slow Injection Method for Reducing Tissue Injury

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The author has nothing to disclose.

One of the critical points in the successful performance of the slow injection method is the preparation of an effective injection system using a powerful syringe pump and a strong pressure transfer tube. A high-pressure system is required to push gel out from the tip of a fine needle. The second critical point is the stabilization of the heart. The beating of the heart against an injection needle advanced into the myocardium can injure the tissue. In this study, an echo-guided injection was conducted to avoid the animals undergoing a second open chest injury and to administer the cell injection in a stabilized heart with the lungs inflated. Moreover, in some applications for larger animals or humans, some injection devices attached onto the heart should be considered as part of the strategic design of the application. For open chest injections into the hearts of small animals, the use of a long, flexible needle is recommended given their higher heart rates.
In this work, the slow injection method significantly increased the surviving cardiomyocyte volume compared to the normal injection method. The normal injection causes cell damage via shear stress15. In contrast, the slow injection method does not cause such stress theoretically because it uses a non-Newtonian solution in addition to the slow injection.
In terms of local fibrosis, the interstitial space around the normally injected surviving cardiomyocytes showed strong and widespread type I collagen deposition. In contrast, the type I collagen signals around the engrafted cardiomyocytes grafted using the slow injection method were much weaker and more limited. This suggests that the slow injection method caused significantly less damage. The slow injection of neonatal cardiomyocytes into the adult myocardium significantly improved the contractile function of the infarcted heart. The histological analyses suggested that grafting the cardiomyocytes using the slow injection method resulted in direct connections and functional coupling with the host cardiomyocytes. This phenomenon explains the mechanism of the functional recovery of the host myocardium. To the best of our knowledge, this is the first report of engrafted neonatal cardiomyocytes with large-scale seamless connections to the host adult cardiomyocytes. The functional connections with the host myocardium via electrical and mechanical coupling may make the engrafted cardiomyocytes mature and allow them to act as functional myocytes that contribute to the host heart function. Long-term physical force interactions between the host and graft cardiomyocytes are crucial for full maturation. Therefore, 2 months might be required after the injection for the functional recovery of the infarcted heart. The time-dependent recovery of the patient&#39;s heart function may be an expected phenomenon in therapeutic applications, and this can be a hallmark of the successful establishment of de novo functional coupling and integration between the host and grafted cardiomyocytes.
The slow injection method can be performed during open chest surgery. In addition, this method can be applied to mice. For future applications in human therapy, we still need to resolve several issues. The injection speed should be optimized by considering the buffer capacity of the interstitial fluid flow in each human target organ. Xeno-free materials, such as human gelatin or biodegradable synthetic materials, should be applied. Clinical GMP-grade slow injection apparatus, such as compact organ-specific disposable tools or a reusable wide-organ applicable apparatus, should be developed.

Cell administration and replacement are promising novel therapeutic strategies for heavily damaged organs. Among these novel therapeutic strategies, cardiac regenerative medicine has attracted considerable attention. However, the inflammation caused by injuries mediates scar formation in several organs1,2,3,4. The human heart consists of approximately 1010 cardiomyocytes; therefore, theoretically5,6, it must be treated with more than 109 cardiomyocytes. Administering a large number of cardiomyocytes via traditional injection methods may lead to significant tissue injuries7. This method provides a novel cell injection method with minimal tissue invasion.
Cell administration into the organ parenchyma requires injection(s). However, a discrepancy exists in that the injection itself may lead to tissue injury. Tissue injury causes local inflammation and incurable scarring in organs and tissues, as well as impaired regenerative ability8,9,10. The mammalian heart has an extremely high propensity to develop scars instead of regenerating because it requires immediate injury repair in order to endure the high blood pressure caused by its continuous pumping function11. Ablation therapy utilizes this high propensity toward scar formation and blocks the circuit likely to undergo scar formation using arrhythmia12. In a previous study, it was observed that the scar tissue isolated the injected cardiomyocytes in the host myocardium. Thus, this represents the next target issue that needs to be overcome to obtain improved therapeutic efficacy in cardiac regenerative medicine.
Tissue interstitial fluid flow plays a vital role in conveying oxygen and nutrients to cells and removing the excreted waste from cells. The physiological speed of interstitial fluid flow in each tissue/organ is different (the range is 0.01-10 &#181;m/s)13. To the best of the author&#39;s knowledge, there are no data regarding the capacity of individual tissues/organs to support extra amounts of fluid without pathological edema; however, this experiment attempts to use a slow injection speed to possibly reduce tissue injury, and the results can be used to determine the practicality of this concept.

<table><tbody><tr><td>18-gauge needle &amp;amp; tuberculin, 1 mL</td><td>Terumo&amp;nbsp;</td><td>NN1838R, SS-01T</td><td></td></tr><tr><td>29-gauge 50 mm-long needle</td><td>Ito Corporation, Tokyo, Japan</td><td>14903 Type-A&amp;nbsp;</td><td></td></tr><tr><td>A copper tube</td><td>General Suppliers</td><td></td><td>outer diameter, 1 mm; inner diameter, 0.3 mm; thickness, 0.35 mm</td></tr><tr><td>Ads Buffer</td><td>Each ingredient was purchased from Fuji Film Wako Chemical Inc., Miyazaki, Japan</td><td></td><td>Hand made, Composition: 116 mM NaCl, 20 mM HEPES, 12.5 mM NaH2PO4, 5.6 mM glucose, 5.4 mM KCl, 0.8 mM MgSO4, pH 7.35</td></tr><tr><td>alpha-MEM</td><td>Fuji Film Wako Chemical Inc., Miyazaki, Japan</td><td>051-07615</td><td></td></tr><tr><td>Anti-collagen type I rabbit polyclonal antibody (H+L)</td><td>Proteintech</td><td>14695-1-AP</td><td>using dilution 1:100</td></tr><tr><td>Anti-Connexin-43 rabbit polyclonal antibody (H+L)</td><td>Sigma Aldrich</td><td>C6219</td><td>using dilution 1:100</td></tr><tr><td>Anti-rabbit IgG (H+L) donley polyclonal antibody-AlexaFluo488&amp;nbsp;</td><td>Thermo Scientific</td><td>A21206</td><td>using dilution 1:300</td></tr><tr><td>blocking solution (Blocking One)</td><td>Nacalai Tesque, Kyoto, Japan</td><td>03953-95&amp;nbsp;</td><td></td></tr><tr><td>collagenase</td><td>Fuji Film Wako Chemical Inc., Miyazaki, Japan</td><td>034-22363</td><td></td></tr><tr><td>confocal laser microscope</td><td>Carl Zeiss Inc., Oberkochen, Germany</td><td>LSM510 META</td><td></td></tr><tr><td>DNase I</td><td>Sigma-Aldrich</td><td>DN25</td><td></td></tr><tr><td>FACS Aria III</td><td>Becton Dickinson, Franklin Lakes, NJ, USA</td><td></td><td></td></tr><tr><td>fetal bovine serum</td><td>BioWest, FL, USA</td><td>S1820-500</td><td></td></tr><tr><td>fine movement device (Micromanipulator)</td><td>Narishige Co., Ltd., Tokyo, Japan</td><td>M-44</td><td></td></tr><tr><td>fluorescence microscope</td><td>&amp;nbsp;Nikon Instruments, Tokyo, Japan</td><td>Eclipse Ti2</td><td></td></tr><tr><td>gelatin from bovine skin</td><td>Sigma-Aldrich</td><td>G9382</td><td>dissolving in PBS (-) to 10%, and autoclaving it</td></tr><tr><td>Neonatal Sprague-Dawley (SD) rats</td><td>Japan SLC Inc., Shizuoka, Japan</td><td></td><td>0&amp;ndash;2 d after birth</td></tr><tr><td>non-adhesive 96-well plates (spheloid plate)</td><td>Sumitomo Bakelite, Tokyo, Japan</td><td>MS-0096S</td><td></td></tr><tr><td>Optimal Cutting Temperature (OCT) Compound&amp;nbsp;</td><td>Sakura Finetek USA, Inc., CA, USA</td><td>Tissue-Tek OCT compound</td><td></td></tr><tr><td>peristaltic pump (for cooling system)</td><td>As One Co., Osaka, Japan</td><td>SMP-23AS</td><td></td></tr><tr><td>PKH26</td><td>Sigma-Aldrich</td><td>PKH26GL</td><td></td></tr><tr><td>Stir Bar, Micro, Magnetic, PTFE, Length x Dia. in mm: 5 x 2</td><td>Chemglass life sciences LLC, NJ, USA</td><td>CG-2003-120</td><td></td></tr><tr><td>syringe&amp;nbsp;</td><td>Ito Corporation, Tokyo, Japan</td><td>MS-N25</td><td></td></tr><tr><td>syringe pump with remote controller</td><td>As One Co., Osaka, Japan</td><td>MR-1, CT-10</td><td></td></tr><tr><td>tetramethylrhodamine methyl ester</td><td>Thermo Fisher Scientific, Waltham, MA, USA</td><td>T668</td><td></td></tr><tr><td>trypsin</td><td>DIFCO, Becton Dickinson, Franklin Lakes, NJ, USA</td><td>215240</td><td></td></tr><tr><td>Tween-20</td><td>Fuji Film Wako Chemical Inc., Miyazaki, Japan</td><td>167-11515</td><td></td></tr><tr><td>veterinarian ointment</td><td>&amp;nbsp;Fujita Pharmaceutical Co., Ltd.</td><td>Hibikusu ointment&amp;nbsp; #WAK-95832</td><td></td></tr><tr><td>Vevo 2100 Imaging System</td><td>Fujifilm VisualSonics, Inc., Toronto, Canada</td><td>Vevo 2100</td><td></td></tr><tr><td>Vevo 2100 Imaging System software version 1.0.0</td><td>Fujifilm VisualSonics, Inc., Toronto, Canada</td><td>Vevo 2100</td><td></td></tr><tr><td>Weakly curved needle with ophthalmic thread</td><td>Natsume Seisakusho Co., Ltd., Tokyo, Japan</td><td>C7-70</td><td></td></tr></tbody></table>

a novel cell injection method with minimum invasion

Directly injecting cells into tissues is a necessary process in cell administration and/or replacement therapy. The cell injection requires a sufficient amount of suspension solution to allow the cells to enter the tissue. The volume of the suspension solution affects the tissue, and this can cause major invasive injury as a result of the cell injection. This paper reports on a novel cell injection method, called slow injection, that aims to avoid this injury. However, pushing out the cells from the needle tip requires a sufficiently high injection speed according to Newton's law of shear force. To solve the above contradiction, a non-Newtonian fluid, such as gelatin solution, was used as the cell suspension solution in this work. Gelatins solution have temperature sensitivity, as their form changes from gel to sol at approximately 20 &#176;C. Therefore, to maintain the cell suspension solution in the gel form, the syringe was kept cooled in this protocol; however, once the solution was injected into the body, the body temperature converted it to a sol. The interstitial tissue fluid flow can absorb excess solution. In this work, the slow injection technique allowed cardiomyocyte balls to enter the host myocardium and engraft without surrounding fibrosis. This study employed a slow injection method to inject purified and ball-formed neonatal rat cardiomyocytes into a remote area of myocardial infarction in the adult rat heart. At 2 months following the injection, the hearts of the transplanted groups showed significantly improved contractile function. Furthermore, histological analyses of the slow-injected hearts revealed seamless connections between the host and graft cardiomyocytes via intercalated disks containing gap junction connections. This method could contribute to next-generation cell therapies, particularly in cardiac regenerative medicine.

Effects of the slow injection on cell survival and collagen deposition 
Neonatal rat cardiomyocyte balls labeled with PKH26 were injected into normal nude rat myocardium using a normal or slow injection method. The results showed that the slow injection method significantly increased the engrafted cell volume (Figure 3A) and significantly decreased on-site type I collagen deposition (Figure 3B).
Effects of slow injection on treatment efficacy in a rat infarction model 
The echocardiograph-guided slow injection method was used to inject neonatal rat cardiomyocyte balls or PBS(&#8722;) into the infarcted hearts of model rats. The cell-injected group alone exhibited significant improvement in heart contraction function after 2 months (Figure 4A). Immunohistochemical analyses revealed a seamless connection between the engrafted cells and host myocytes via intercalated disks containing gap junctions (Figure 4B).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65260/65260fig01.jpg" /> 
Figure 1: Schematic of the whole injection system. (A) Main injection apparatus. (B) Injection syringe cooling system. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65260/65260fig01large.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/65260/65260fig02.jpg" /> 
Figure 2: Echocardiography-guided percutaneous slow injection. (A) Setting of the animal, echo probe, and injection apparatus. (B) Echocardiographic view of the injecting syringe and heart. Note that the left and the right pictures are the same, but a yellow line has been added to indicate the needle position. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65260/65260fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65260/65260fig03.jpg" /> 
Figure 3: Effect of the slow injection method on the engrafted cell volume and collagen deposition. (A) The engrafted cell volumes (N = 3) were calculated from serial sections. The error bars indicate standard deviations. *P &#60; 0.01 in a non-paired t-test. (B) Immunohistochemical staining for type I collagen. The scale bar indicates 200 &#181;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65260/65260fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65260/65260fig04.jpg" /> 
Figure 4: Improvements in heart function and histological integration with cardiomyocyte balls engrafted by the slow injection method. (A) Representative echocardiograph M-mode views. The graph shows the transition of fraction shortenings in the cardiomyocyte ball-transplanted group (solid red line; N = 4) and vehicle (cell suspension solution for the slow injection method) group (dotted blue line; N = 3). Abbreviations: MI = myocardial infarction; Cx43 = connexin 43; DAPI = 4&#39;,6-diamidino-2-phenylindole; PKH26 = red fluorescent cell membrane label. The error bars indicate the standard deviations. *P &#60; 0.01 in a paired t-test. (B) Immunohistochemical analyses of the relationship between the engrafted cardiomyocyte balls and host cardiomyocytes. The conventional laser microscopic observations using a 2x objective lens are shown in the left column. Zoomed-in versions (using a 20x objective lens) of two areas shown in the box, labeled with * and #, are presented below. Scale bars: top image = 300 &#181;m; * and # = 30 &#181;m. Confocal laser microscopic images using a 20x objective lens are shown along for comaparison. Three positions are shown. In the merged images, the arrowheads indicate the existence of gap junctions (Cx43) directly connecting the graft and host cardiomyocytes. The scale bar indicates 30 &#181;m. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65260/65260fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Video 1: Echo-guided slow injection method. The B-mode echocardiogram in the frontal view shows the injection needle tip advanced into the myocardium. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65260/Supplementary video.mp4" target="_blank">Please click here to download this File.</a>

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Author Spotlight: A Novel Cell Injection Method with Minimum Invasion  

This method eliminates any major invasion during cell injections caused by the cell suspension solution.

A Novel Cell Injection Method with Minimum Invasion

Directly injecting cells into tissues is a necessary process in cell administration and/or replacement therapy. The cell injection requires a sufficient ...

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Research

JoVE Journal

Bioengineering

1.3K Views.  Kansai Medical University. This method eliminates any major invasion during cell injections caused by the cell suspension solution.

Video: Author Spotlight: A Novel Cell Injection Method with Minimum Invasion  

Intra-iliac Artery Injection for Efficient and Selective Modeling of Microscopic Bone Metastasis

Intra-iliac artery (IIA) injection is an efficient approach to introduce metastatic lesions of various cancer cells in animals. Compared to the widely used intra-cardiac and intra-tibial injections, IIA injection brings several advantages. First, it can deliver a large quantity of cancer cells specifically to hind limb bones, thereby providing spatiotemporally synchronized early-stage colonization events and allowing robust quantification and swift detection of disseminated tumor cells. Second, it injects cancer cells into the circulation without damaging the local tissues, thereby avoiding inflammatory and wound-healing processes that confound the bone colonization process. Third, IIA injection causes very little metastatic growth in non-bone organs, thereby preventing animals from succumbing to other vital metastases, and allowing continuous monitoring of indolent bone lesions. These advantages are especially useful for the inspection of progression from single cancer cells to multi-cell micrometastases, which has largely been elusive in the past. When combined with cutting-edge approaches of biological imaging and bone histology, IIA injection can be applied to various research purposes related to bone metastases.

This manuscript provides the detailed procedure of intra-iliac artery (IIA) injection, a technique to deliver cancer cells specifically to hind limb tissues including bones to establish experimental bone metastases. Although initially established with breast tumor models, this protocol can be easily extended to other cancer types.

Intra-iliac artery (IIA) injection is an efficient approach to introduce metastatic lesions of various cancer cells in animals. Compared to the widely ...

Cancer Research

Intraspinal Cell Transplantation for Targeting Cervical Ventral Horn in Amyotrophic Lateral Sclerosis and Traumatic Spinal Cord Injury

Respiratory compromise due to phrenic motor neuron loss is a debilitating consequence of a large proportion of human traumatic spinal cord injury (SCI) cases 1 and is the ultimate cause of death in patients with the motor neuron disorder, amyotrophic laterals sclerosis (ALS) 2.
ALS is a devastating neurological disorder that is characterized by relatively rapid degeneration of upper and lower motor neurons. Patients ultimately succumb to the disease on average 2-5 years following diagnosis because of respiratory paralysis due to loss of phrenic motor neuron innnervation of the diaphragm 3. The vast majority of cases are sporadic, while 10% are of the familial form. Approximately twenty percent of familial cases are linked to various point mutations in the Cu/Zn superoxide dismutase 1 (SOD1) gene on chromosome 21 4. Transgenic mice 4,5 and rats 6 carrying mutant human SOD1 genes (G93A, G37R, G86R, G85R) have been generated, and, despite the existence of other animal models of motor neuron loss, are currently the most highly used models of the disease.
Spinal cord injury (SCI) is a heterogeneous set of conditions resulting from physical trauma to the spinal cord, with functional outcome varying according to the type, location and severity of the injury 7. Nevertheless, approximately half of human SCI cases affect cervical regions, resulting in debilitating respiratory dysfunction due to phrenic motor neuron loss and injury to descending bulbospinal respiratory axons 1. A number of animal models of SCI have been developed, with the most commonly used and clinically-relevant being the contusion 8.
Transplantation of various classes of neural precursor cells (NPCs) is a promising therapeutic strategy for treatment of traumatic CNS injuries and neurodegeneration, including ALS and SCI, because of the ability to replace lost or dysfunctional CNS cell types, provide neuroprotection, and deliver gene factors of interest 9.
Animal models of both ALS and SCI can model many clinically-relevant aspects of these diseases, including phrenic motor neuron loss and consequent respiratory compromise 10,11. In order to evaluate the efficacy of NPC-based strategies on respiratory function in these animal models of ALS and SCI, cellular interventions must be specifically directed to regions containing therapeutically relevant targets such as phrenic motor neurons. We provide a detailed protocol for multi-segmental, intraspinal transplantation of NPCs into the cervical spinal cord ventral gray matter of neurodegenerative models such as SOD1G93A mice and rats, as well as spinal cord injured rats and mice 11.

Neural precursor transplantation is a promising strategy for protecting and/or replacing lost/dysfunctional cervical phrenic motor neurons in spinal cord injury (SCI) and the motor neuron disorder, amyotrophic laterals sclerosis (ALS). We provide a protocol for cell delivery to cervical spinal cord ventral horn in rodent models of ALS and SCI.

Respiratory compromise due to phrenic motor neuron loss is a debilitating consequence of a large proportion of human traumatic spinal cord injury (SCI) ...

Medicine

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 ...

Minimally-invasive Technique for Injection into Rat Optic Nerve

The rat optic nerve is a useful model for stem cell regeneration research. Direct injection into the rat optic nerve allows delivery into the central nervous system in a minimally-invasive surgery without bone removal. This technique describes an approach to visualization and direct injection of the optic nerve following minor fascial dissection from the orbital ridge, using a conjunctival traction suture to gently pull the eye down and out. Representative examples of an injected optic nerve show successful injection of dyed beads.

Direct injection into the rat optic nerve is useful for regenerative research. We demonstrate a minimally-invasive technique for direct injection into a rat optic nerve that does not involve opening the skull. Using this method, surgical complications are minimized and recovery is more rapid.

The rat optic nerve is a useful model for stem cell regeneration research. Direct injection into the rat optic nerve allows delivery into the central ...

Neuroscience

An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Subretinal injections have been successfully used in both humans and rodents to deliver therapeutic interventions of proteins, viral agents, and cells to the interphotoreceptor/subretinal compartment that has direct exposure to photoreceptors and the retinal pigment epithelium (RPE). Subretinal injections of plasminogen as well as recent preclinical and clinical trials have demonstrated safety and/or efficacy of delivering viral vectors and stem cells to individuals with advanced retinal disease. Mouse models of retinal disease, particularly hereditary retinal dystrophies, are essential for testing these therapies. The most common injection procedure in rodents is to use small transcorneal or transcleral incisions with an anterior approach to the retina. With this approach, the injection needle penetrates the neurosensory retina disrupting the underlying RPE and on insertion can easily nick the lens, causing lens opacification and impairment of noninvasive imaging. Accessing the subretinal space via a transcleral, posterior approach avoids these problems: the needle crosses the sclera approximately 0.5 mm from the optic nerve, without retinal penetration and avoids disrupting the vitreous. Collateral damage is limited to that associated with the focal sclerotomy and the effects of a transient, serous retinal detachment. The simplicity of the method minimizes ocular injury, ensures rapid retinal reattachment and recovery, and has a low failure rate. The minimal damage to the retina and RPE allows for clear assessment of the efficacy and direct effects of the therapeutic agents themselves. This manuscript describes a novel subretinal injection technique that can be used to target viral vectors, pharmacological agents, stem cells or induced pluripotent stem (iPS) cells to the subretinal space in mice with high efficacy, minimal damage, and fast recovery.

An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Subretinal injections are the most common technique for delivering large therapeutic agents such as proteins and viral vectors to photoreceptors and the retinal pigment epithelium. An alternative method in mice that successfully targets the subretinal space with minimal collateral damage and fast recovery times is described here.

Subretinal injections have been successfully used in both humans and rodents to deliver therapeutic interventions of proteins, viral agents, and cells to ...

Manipulation of Single Neural Stem Cells and Neurons in Brain Slices using Robotic Microinjection

A central question in developmental neurobiology is how neural stem and progenitor cells form the brain. To answer this question, one needs to label, manipulate, and follow single cells in the brain tissue with high resolution over time. This task is extremely challenging due to the complexity of tissues in the brain. We have recently developed a robot, that guide a microinjection needle into brain tissue upon utilizing images acquired from a microscope to deliver femtoliter volumes of solution into single cells. The robotic operation increases resulting an overall yield that is an order of magnitude greater than manual microinjection and allows for precise labeling and flexible manipulation of single cells in living tissue. With this, one can microinject hundreds of cells within a single organotypic slice. This article demonstrates the use of the microinjection robot for automated microinjection of neural progenitor cells and neurons in the brain tissue slices. More broadly, it can be used on any epithelial tissue featuring a surface that can be reached by the pipette. Once set up, the microinjection robot can execute 15 or more microinjections per minute. The microinjection robot because of its throughput and versality will make microinjection a broadly straightforward high-performance cell manipulation technique to be used in bioengineering, biotechnology, and biophysics for performing single-cell analyses in organotypic brain slices.

This protocol demonstrates the use of a robotic platform for microinjection into single neural stem cells and neurons in brain slices. This technique is versatile and offers a method of tracking cells in tissue with high spatial resolution.

A central question in developmental neurobiology is how neural stem and progenitor cells form the brain. To answer this question, one needs to label, ...

Injection of Porcine Adipose Tissue-Derived Stroma Cells via Waterjet Technology

Urinary incontinence (UI) is a highly prevalent condition characterized by the deficiency of the urethral sphincter muscle. Regenerative medicine branches, particularly cell therapy, are novel approaches to improve and restore the urethral sphincter function. Even though injection of active functional cells is routinely performed in clinical settings by needle and syringe, these approaches have significant disadvantages and limitations. In this context, needle-free waterjet (WJ) technology is a feasible and innovative method that can inject viable cells by visual guided cystoscopy in the urethral sphincter. In the present study, we used WJ to deliver porcine adipose tissue-derived stromal cells (pADSCs) into cadaveric urethral tissue and subsequently investigated the effect of WJ delivery on cell yield and viability. We also assessed the biomechanical features (i.e., elasticity) by atomic force microscopy (AFM) measurements. We showed that WJ delivered pADSCs were significantly reduced in their cellular elasticity. The viability was significantly lower compared to controls but is still above 80%.

We present a method of cell injection via needle free waterjet technology coupled with a sequela of post-delivery investigations in terms of cellular viability, proliferation, and elasticity measurements.

Urinary incontinence (UI) is a highly prevalent condition characterized by the deficiency of the urethral sphincter muscle. Regenerative medicine ...

Single-cell Microinjection for Cell Communication Analysis

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a hemichannel by each neighboring cell to form the gap junction. In mammalian cells, the hemichannel is formed by six connexins, monomers with four transmembrane domains and a C and N terminal within the cytoplasm. Gap junctions permit the exchange of ions, second messengers, and small metabolites. In addition, they have important roles in many forms of cellular communication within physiological processes such as synaptic transmission, heart contraction, cell growth and differentiation. We detail how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells. It is expected that in functional gap junctions, the dye will diffuse from the loaded cell to the connected cells. It is a very useful technique to study gap junctions since you can evaluate the diffusion of the fluorescence in real time. We discuss how to prepare the cells and the micropipette, how to use a micromanipulator and inject a low molecular weight fluorescent dye in an epithelial cell line.

We describe here how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells, and provide some useful tips. We expect that this paper will help everyone to evaluate the degree of cellular coupling due to functional gap junctions. Everything described here could be, in principle, adapted to other fluorescent dyes with molecular weight below 1,000 Daltons.

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a ...

Biology

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter implantation. The technical limitation of this surgical technique lies in the finesse of the hands. The injection is rapid, especially for a trained experimenter, and since tissue disruption with this technique is minimal, repeated injections are possible; moreover immune reaction to foreign tools (e.g. catheter) does not occur, thereby giving a better and more specific read out of spinal cord modulation. Since the application of the substance is largely limited to the target region of the spinal cord, drugs do not need to be applied in large dosages, and more importantly unwanted effects on other tissue, as observed with a systemic delivery, could be circumvented1,2. Moreover, we combine this technique with in vivo transfection of nucleic acid with the help of polyethylenimine (PEI) reagent3, which provides tremendous versatility for studying spinal functions via delivery of pharmacological agents as well as gene, RNA, and protein modulators.

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a simple and rapid technique of intrathecal needle puncture for a localized transfection of siRNA in the lumbar spinal cord in mouse under short lasting light anesthesia.

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter ...

A Minimally Invasive, Accurate, and Efficient Technique for Intrathymic Injection in Mice

Intrathymic injection in mouse models is an important technique for studying thymic and immune function, including genetic and acquired T cell disorders. This requires methods for the direct deposition of reagents and/or cells into the thymus of living mice. Traditional methods of intrathymic injection include thoracic surgery or minimally invasive percutaneous blind injections, both of which have significant limitations. Ultra-high frequency ultrasound imaging devices have made image-guided percutaneous injections possible in mice, greatly improving the injection accuracy of the percutaneous injection approach and enabling the injection of smaller targets. However, image-guided injections rely on the utilization of an integrated rail system, making this a rigid and time-consuming procedure. A unique, safe, and efficient method for percutaneous intrathymic injections in mice is presented here, eliminating reliance on the rail system for injections. The technique relies on using a high-resolution micro-ultrasound unit to image the mouse thymus noninvasively. Using a free-hand technique, a radiologist can place a needle tip directly into the mouse thymus under sonographic guidance. Mice are cleaned and anesthetized before imaging. For an experienced radiologist adept at ultrasound-guided procedures, the learning period for the stated technique is quite short, typically within one session. The method has a low morbidity and mortality rate for the mice and is much faster than current mechanically assisted techniques for percutaneous injection. It allows the investigator to efficiently perform precise and reliable percutaneous injections of thymuses of any size (including very small organs such as the thymus of aged or immunodeficient mice) with minimal stress on the animal. This method enables the injection of individual lobes if desired and facilitates large-scale experiments due to the time-saving nature of the procedure.

The present protocol describes an interventional radiology procedure established for intrathymic injection in mice to avoid the risk of open surgery and improve the accuracy of blind percutaneous injections.

Intrathymic injection in mouse models is an important technique for studying thymic and immune function, including genetic and acquired T cell disorders. ...

A Novel Cell Injection Method with Minimum Invasion

Summary

Explore More Videos

Chapters in this video

Intra-iliac Artery Injection for Efficient and Selective Modeling of Microscopic Bone Metastasis

Intraspinal Cell Transplantation for Targeting Cervical Ventral Horn in Amyotrophic Lateral Sclerosis and Traumatic Spinal Cord Injury

Intramyocardial Cell Delivery: Observations in Murine Hearts

Minimally-invasive Technique for Injection into Rat Optic Nerve

An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Manipulation of Single Neural Stem Cells and Neurons in Brain Slices using Robotic Microinjection

Injection of Porcine Adipose Tissue-Derived Stroma Cells via Waterjet Technology

Single-cell Microinjection for Cell Communication Analysis

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

A Minimally Invasive, Accurate, and Efficient Technique for Intrathymic Injection in Mice