We thank our co-authors from the original publications for their help and support.

The porcine adipose tissue samples were obtained from the Institute for Experimental Surgery at the University of Tuebingen. All procedures were approved by local animal welfare authorities under the animal experiment number CU1/16.
1. Isolation of porcine adipose tissue-derived stromal cells
<ol>
	<li>Use porcine adipose tissue delivered from the Institute for Experimental Surgery in a 50 mL centrifuge tube to the laboratory.</li>
	<li>Transfer the tissue to a sterile Petri dish under the sterile bench and mince it with two scalpels (No. 10) to small fragments and mush. 
	NOTE: Scissors can also be used in order to get small fragments. The smaller the fragments are, the better for the following digestion.
	<ol>
		<li>Transfer the small fragments/mush into a 50 mL centrifuge tube and incubate it with 5 mL of homogenizer solution for 30 min at 37 &#176;C on a shaker. The homogenizer solution is PBS with 1% bovine serum albumin (BSA) (stock solution: 1 % (w/v) BSA in PBS) and 0.1% collagenase type I. 
		NOTE: Always prepare homogenizer solution freshly. The shaker has a circular shaking motion at a slow speed 25-500 rpm.</li>
	</ol>
	</li>
	<li>To stop the incubation, add 10 mL of growth media [Dulbecco&#39;s Modified Eagle&#39;s Medium - low glucose (DMEM-LG), 2.5% HEPES sodium salt solution (1 M), 10% fetal bovine serum (FBS), 1% L-Glutamin (200 mM), 1% Penicillin-Streptomycin (10000 U/mL Penicillin; 10,000 &#181;g/mL Streptomycin) and 1% Amphotericin B (250 &#181;g/mL)]. 
	NOTE: Growth media can be prepared beforehand. Antibiotics and antifungal drugs are necessary in the preparation of the growth media for cell culture to protect cells from contaminations.</li>
	<li>Incubate the centrifuge tubes for 10 min at room temperature.</li>
	<li>Aspirate the fraction with adipocytes, which is lighter and therefore swimming on top of the liquid, with a 10 mL pipette and discard it.</li>
	<li>Filter the remaining stromal fraction through a 100 &#181;m cell sieve with a polyethylene terephthalate (PET) membrane free of heavy metals to withhold adipose tissue fragments and to retrieve only the cell suspension.</li>
	<li>Centrifuge the filtered cell suspension for 7 min at 630 x g at room temperature.</li>
	<li>Resuspend the cell pellet in 2 mL of PBS to wash the cells once.</li>
	<li>Centrifuge the cell suspension again for 7 min at 630 x g at room temperature.</li>
	<li>After centrifugation and repeated resuspension of the cell pellet in 10 mL of growth media per flask, seed cells in 75 cm2 cell culture flasks. 
	&#8203;NOTE: Depending on the size of the cell pellet after the washing step with PBS, seed cells in one to four flasks.</li>
</ol>
2. Cell cultivation of porcine adipose tissue-derived stromal cells
<ol>
	<li>Harvest and passage cells when they reach 70% confluence.</li>
	<li>Wash cells with 10 mL of PBS twice and aspirate PBS completely. 
	NOTE: This step is necessary to remove remaining growth media, which is complemented by 10% FBS. FBS has several protease inhibitors which can hinder the function of the following trypsinization.</li>
	<li>Add 3 mL of 0.05%-Trypsin-EDTA per flask to detach the cells.</li>
	<li>Incubate flasks for 3 min at 37 &#176;C.</li>
	<li>Check under the microscope if cells are detached. 
	NOTE: Sometimes it takes some more time to detach the cells. In this case, incubate flasks for maximum 5 min at 37 &#176;C.</li>
	<li>To stop the incubation and detaching process, add 3 mL of growth media. 
	NOTE: As noted above, growth media is complemented by 10% FBS that contains protease inhibitors.</li>
	<li>Transfer the detached cells to a centrifugation tube and centrifuge for 7 min at 630 x g at room temperature.</li>
	<li>Resuspend the cells in 10 mL of growth media and count them with a hemocytometer.</li>
	<li>Seed cells with an inoculation density of 3 x 105 cells per 75 cm2 flask.</li>
</ol>
3. Labelling of cells with calcein-AM
NOTE: Cells that are injected into cadaveric tissues are stained with a green-fluorescent membrane-permeable live-cell stain and a red-fluorescent membrane-impermeant viability indicator to verify that extracted cells are the same as the injected cells and not tissue fragments of the urethra.
<ol>
	<li>Wash cells twice with 10 mL of PBS.</li>
	<li>Add 5 mL of dye solution per 75 cm2 flask. The dye solution consists of growth media with 2 &#181;M of the green-fluorescent membrane-permeable live-cell dye and 4 &#181;M red-fluorescent membrane-impermeant viability indicator.</li>
	<li>Incubate for 30 min at room temperature in the dark.</li>
	<li>Aspirate and discard the dye solution.</li>
	<li>Wash cells again twice with 10 mL of PBS.</li>
	<li>Add growth media and document the green and red fluorescence signal by fluorescence microscopy.
	<ol>
		<li>Place the 75 cm2 flask on the microscope table, use the 10x objective, select no fluorescence filter and make sure that the transmitted light path is active. 
		NOTE: These steps are all performed manually on the microscope.</li>
		<li>In parallel to step 3.6.1, open the software program.</li>
		<li>Press the Live button under the Locate tab to obtain a live image of the cells in the flask on the table and focus on them with the coarse and fine adjustment drives. 
		NOTE: On the right sight the live image will appear once the Live button is activated.</li>
		<li>In the sub-tab Microscope Components, select the correct objective (10x).</li>
		<li>In the sub-tab Camera, apply a checkmark for Auto Exposure.</li>
		<li>Press the button Snap to take the first picture with transmitted light.</li>
		<li>Insert the scale bar by using the Graphics tab in the menu bar and selecting Scale Bar.</li>
		<li>Save the image by using the Files tab in the menu bar and selecting Save as czi.</li>
		<li>For the fluorescence pictures change the filter to the one specific for green or red fluorescence and select the reflected light path. 
		NOTE: These changes have to be done manually on the microscope.</li>
		<li>Press again the Live button under the Locate tab to obtain a live image of the cells.</li>
		<li>Press the button Snap to take the second picture with either green or red fluorescence.</li>
		<li>Insert the scale bar by using the Graphics tab in the menu bar and selecting Scale Bar.</li>
		<li>Save the image by using the Files tab in the menu bar and selecting Save as czi.</li>
		<li>Repeat steps 3.6.9 to 3.6.13 with the other fluorescence channel.</li>
	</ol>
	</li>
</ol>
4. Prepare urethral tissue samples for injections
<ol>
	<li>Dissect the urethra and the connecting bladder out of the pig. 
	NOTE: Urethral tissue samples from fresh cadaveric samples of adult female landrace pigs were used for the experiments.</li>
	<li>Transport it to the laboratory in bags on wet ice. 
	NOTE: The samples should be left on ice until further processing. No additives were added to the samples.</li>
	<li>Place the urethra with the bladder on a sponge, which mimics the elasticity of the lower pelvic floor.
	<ol>
		<li>Use the bladder to determine the orientation (proximal, distal, dorsal, and ventral) of the urethra. This is possible due to the localization of the ureters and the three ligaments which fix the bladder in the abdominal and pelvic cavity. The three ligaments are the two vesicae lateralia ligaments on the sides of the bladder and the vesicae medianum, which arises from the ventral surface of the bladder (Figure 1).</li>
	</ol>
	</li>
	<li>Cut open the urethra longitudinally on the dorsal side with aid of a catheter.</li>
	<li>Inject the cells into the opened urethra as described in the following chapters.</li>
</ol>
5. Injections of cells via a Williams needle in fluids and tissue samples
<ol>
	<li>Harvest cells as described in step 2.</li>
	<li>In contrast to step 2.9, adjust cell density for injections to 2.4 x 106 cells per mL. 
	NOTE: For injections into tissue samples label the cells with a green-fluorescent membrane-permeable live-cell.</li>
	<li>Aspirate the cell suspension with a syringe and apply the Williams cystoscopic injection needle (WN) to it.</li>
	<li>Either hold the needle shortly above the 2 mL of growth media in a 15 mL centrifugation tube or insert the needle into the opened urethra tissue. In both cases inject 250 &#181;L of cells manually. 
	NOTE: Cells injected into the cadaveric urethra form an injection dome.</li>
	<li>Collect cells injected into media directly by centrifugation for 7 min at 630 x g at room temperature.</li>
	<li>For cells injected into the cadaveric urethra, aspirate them out of the injection dome with an 18G needle applied on a syringe.</li>
	<li>Transfer the injected and aspirated cells into a centrifugation tube and centrifuge for 7 min at 630 x g at room temperature comparably to the cells injected into media.</li>
	<li>After centrifugation, resuspend the cells in 4 mL of growth media in both cases.</li>
	<li>Determine cell yield and viability by aid of Trypan blue dye exclusion with a hemocytometer.
	<ol>
		<li>Mix 20 &#181;L of cell suspension thoroughly with 20 &#181;L of Trypan blue.</li>
		<li>Fill 10 &#181;L of this mixture in each chamber of the hemocytometer. 
		NOTE: Both chambers are filled and counted to gain statistical optimization by doubling sampling.</li>
		<li>Under a microscope, count cells in all four corner squares. 
		NOTE: Count cells shining in white (unstained) as viable cells whereas cells shining blue (stained) are counted as dead cells.</li>
		<li>To calculate the cell number per mL, calculate the mean of the two chambers, divide this number by two and multiply it with 104.</li>
	</ol>
	</li>
</ol>
6. Injections of cells via Waterjet in fluids and tissue samples
<ol>
	<li>Harvest cells as described in step 2.</li>
	<li>In contrast to steps 2.9 and 5.2, adjust cell density for injections to 6 x 106 cells per mL. 
	NOTE: For injections into tissue samples use cells labelled with a green-fluorescent membrane-permeable live-cell.</li>
	<li>Fill the cell suspension into the dosing unit of the WJ device.</li>
	<li>Either hold the injection nozzle shortly above the 2 mL of growth media in a 15 mL centrifugation tube or shortly above the opened urethra tissue.</li>
	<li>In both cases inject 100 &#181;L of cells by the device using a high pressure for tissue penetration followed by a low-pressure phase for cell injections. Use the pressure settings E60-10. 
	NOTE: Cells injected into the cadaveric urethra form an injection dome.</li>
	<li>Collect cells injected into media directly by centrifugation for 7 min at 630 x g at room temperature.</li>
	<li>For cells injected into the cadaveric urethra, aspirate them out of the injection dome with an 18G needle applied on a syringe.</li>
	<li>Transfer the injected and aspirated cells into a centrifugation tube and centrifuge for 7 min at 630 x g at room temperature comparably to the cells injected into media.</li>
	<li>After centrifugation, resuspend cells in 4 mL of growth media in both cases.</li>
	<li>Determine cell yield and viability by aid of Trypan blue dye exclusion with a hemocytometer as described in detail in 5.10.</li>
</ol>
7. Biomechanical assessment of cellular elasticity by atomic force microscopy (AFM)
<ol>
	<li>Preparation of samples 
	NOTE: The retrieved cells following injection are now subject for further analyses by AFM. Additionally, cells that were not injected are used as controls.
	<ol>
		<li>Seed cells in tissue culture dishes with a density of 5 x 105 cells per dish. 
		NOTE: Seed one tissue culture dish per condition. The conditions are controls, ADSCs injected by WJ or WN into media and ADSCs injected by WJ or WN into cadaveric tissue.</li>
		<li>Incubate cells in tissue culture dishes for 3 h at 37 &#176;C. 
		NOTE: To avoid variances due to cells in G1 phase and during mitosis, we performed AFM measurements 3 h after the cell seeding step.</li>
		<li>Right before the measurement with the AFM replace the growth media with 3 mL of Leibovitz&#39;s L-15 media without L-glutamine.</li>
		<li>Place the tissue culture dish in the sample holder of the AFM device and turn on the Petri dish heater set at 37 &#176;C.</li>
	</ol>
	</li>
	<li>Preparation of AFM and cantilever calibration
	<ol>
		<li>Use a glass block that is specified for measurements in liquids and adjust it on the AFM holder. 
		NOTE: The upper surface of the glass block must be straight and in parallel to the AFM holder.</li>
		<li>Carefully place the cantilever on the surface of the glass block. The tip A must lay over the polished optical plane. 
		NOTE: Do not scratch the polished optical surface of the glass block because scratches could lead to interferences in the measurements. The tip A must protrude over the polished optical plane because otherwise the reflection of the laser of the AFM to the photodetector is not possible.</li>
		<li>To stabilize the cantilever on the glass block, add a metallic spring with the aid of tweezers.</li>
		<li>When the cantilever is fixed on the glass block place the block on the AFM head and lock the integrated locking mechanism.</li>
		<li>Mount the AFM head on the AFM device. 
		NOTE: The spring must face the left side to be placed correctly. If this is not the case, correct the position of the glass block and adjust the AFM head again.</li>
		<li>Initialize the software setup and open the software together with the laser alignment window and the approach parameter settings. Additionally, turn on the stepper motor, the laser light and the CCD-camera.</li>
		<li>Identify the cantilever tip A by using the CCD-camera.</li>
		<li>Lower the cantilever until it is fully dipped into the medium. Use the stepper motor function to reach this aim.</li>
		<li>Once the cantilever is fully covered by medium, the laser is aligned on top of the cantilever by using the adjustment screws. The reflected beam must fall onto the center of the photodetector and the sum of the signals must be 1 V or higher. The lateral and vertical deflections should be close to 0. 
		NOTE: If the center is reached can be monitored in the laser alignment function as well as the signal values. If the signal values are not correct, further adjustments are necessary.</li>
		<li>Run now the scanner Approach with the approach parameters (Table 1).</li>
		<li>Retract the cantilever by 100 &#181;m once it reaches the bottom by pressing the Retract button. 
		NOTE: Make sure that no cell is attached at that field were the approach is done because this would falsify the calibration.</li>
		<li>Set up the Run parameters according to the following specifications: Setpoint 1 V, Pulling Length 90 &#181;m, Velocity: 5 &#181;m/s, Sample rate: 2000 Hz and as a Delay Mode: Constant Force.</li>
		<li>Start the calibration force-distance curve measuring by pressing the button Run. 
		NOTE: By clicking the Run button in the software a force-distance curve is obtained.</li>
		<li>Select the region of linear fit of the retracted curve in the software on the obtained calibration force-distance curve. Calculate the spring constant measurement by the software.</li>
		<li>Calibrate the cantilever following the exact parameters and steps as described by Danalache et al.17.</li>
	</ol>
	</li>
	<li>Measuring the elasticity of individual cells
	<ol>
		<li>Visually identify a cell and focus on it. Place the computer mouse at the middle of the cell. To improve the measurements precision, position the AFM tip directly above the cell nucleus. 
		NOTE: The computer mouse is used as visual marker to identify the target site of indentation.</li>
		<li>Now focus on the cantilever and move it on the computer mouse.</li>
		<li>Start the measurement with Run with the parameters given in Table 2. 
		NOTE: The set point parameter obtained by calibration of the cantilever are used. Furthermore, one cell is measured three times. Measure at least 50 cells.</li>
	</ol>
	</li>
	<li>Data processing
	<ol>
		<li>Process the data following exact steps and parameters as previously described by Danalache17.</li>
		<li>Save and export the file.</li>
	</ol>
	</li>
</ol>
8. Statistical analysis
<ol>
	<li>Open the statistical software.</li>
	<li>Choose the selection of New Dataset. 
	NOTE: Two files are opened. One is the &#34;DataSet&#34; and the other one is the &#34;Output&#34; file.</li>
	<li>Select the DataSet file and open the Variable View tab.</li>
	<li>Insert the numeric variables for site injection (capture fluid or cadaveric tissue), category (control, WJ injection, WN injection) and elasticity.</li>
	<li>Insert the measured elasticity data with their corresponding site injection and category number in the Data View tab.</li>
	<li>Analyze the data by selecting the menu bar tab Analyze | Descriptive Statistics and choose Exploratory data analysis.</li>
	<li>As the dependent variable, select Elasticity and factor list select Category. 
	NOTE: The results are displayed in the &#34;Output&#34; file including a box plot which is used for the results section.</li>
	<li>To perform a statistical test, select the Independent Samples within the Nonparametric Test under the menu bar tab Analyze.</li>
	<li>In the new opened file keep the settings in the tab Objective and Settings.</li>
	<li>Open the tab Fields and choose Elasticity as Test Fields and Category as Groups.</li>
	<li>Press Run. 
	NOTE: The results are displayed in the &#34;Output&#34; file. For these analyzes a Mann-U-Whitney test is performed.</li>
	<li>Include the result of the nonparametric test in the box plot of the exploratory data analysis.</li>
	<li>Save the files by selecting File in the menu bar and choosing Save.</li>
</ol>

The authors J.K., M.D, T.A., A.S., W.K. A. have nothing to disclose. The authors W.L. and M.D.E. are employees of ERBE Medizintechnik Lt. T&#252;bingen, the producer of the ERBEJet2 and the WJ prototype employed in this study.

In the present study, we demonstrated and presented a step-by-step approach for WJ cell delivery procedure and employed a sequela of quantitative investigations to assess the effect of WJ delivery on cellular characteristics: cellular viability and biomechanical features (i.e., EM). Following WJ injection, 85.9% of the harvested cells were viable. In terms of WN injection, 97.2% of the cells retained their viability after injection. Thus, the WJ approach fulfills an absolute requirement for a clinical implementation: mor...

Urinary incontinence (UI) is a widespread disorder with a prevalence of 1.8 - 30.5% in European populations1 and is characterized primarily by malfunctioning of the urethral sphincter. From a clinical perspective, surgical treatment is often offered to patients when conservative therapies or physiotherapy fail to address and alleviate the emerging symptoms.
Cell therapy for the potential regenerative repair of the sphincter complex malfunction has been emerging as an avant-garde approach for the treatment of UI pathology2,3. Its main goals are to replace, repair and restore the biological functionality of the damaged tissue. In animal models for UI, stem cell transplantation has shown promising results in urodynamic outcomes2,4,5. Stem cells arise as optimal cellular candidates as they have the ability to undergo self-renewal and multipotent differentiation, thus, aiding the affected tissue regeneration6. Despite the forthcoming regenerative potential, the practical use of cell therapy remains hindered as minimally invasive delivery of cells still face several challenges concerning the injection precision and coverage of the target. Even though the current approach used for cell delivery is injection through a needle-syringe system7, it usually results in an overall deficit of viable cells, with reported viabilities as low as 1%- 31% post-transplantation8. In addition, cell delivery via needle injection has been also shown to affect the placement, the retention rate, as well as distribution of transplanted cells into the targeted tissue9,10,11. A feasible, novel approach that overcomes the abovementioned limitation is the needle-free cell delivery via water-jet technology.
Waterjet (WJ) technology is emerging as a new approach that enables high throughput delivery of cells by cystoscope under visual control in the urethral sphincter12,13. The WJ enables cell delivery at different pressures (E = effects in bar) ranging from E5 to E8013. In the first phase, (tissue penetration phase) isotonic solution is applied with high pressure (i.e., E60 or E80) in order to loosen the extracellular matrix surrounding the tissue targeted and open small interconnecting micro-lacunae. In the second phase (the injection phase), pressure is lowered within milliseconds (i.e., up to E10) in order to gently deliver the cells into the targeted tissue. Following this two step-phase application, the cells are not subjected to additional pressure against the tissue when ejected but are floating in a low-pressure stream into a liquid-filled cavernous area13. In an ex vivo model setting where stem cells were injected via WJ into cadaveric urethra tissue, viable cells could be afterwards aspirated and retrieved from the tissue and further expanded in vitro13. Though a 2020 study by Weber et al. demonstrated the feasibility and applicability of WJ to deliver footprint-free cardiomyocytes into the myocardium14, it has to be borne in mind the WJ technology is still in a prototype stage.
The following protocol describes how to prepare and label porcine adipose tissue-derived stromal cells (pADSC) and how to deliver them into capture fluid and cadaveric tissue via WJ&#160;technology and Williams cystoscopy needles (WN). Post cellular injection, the cellular vitality and elasticity via atomic force microscopy (AFM) is assessed. Via step-by-step instructions, the protocol gives a clear and concise approach to acquire reliable data. The discussion section presents and describes the major advantages, limitations and future perspectives of the technique. The WJ delivery of cells as well as the sequela post translation analyses reported here are replacing the standard needle injection and provide a solid cell delivery framework for regenerative healing of the target tissue. In our recent studies we provided evidence that WJ delivered cells more precisely and at least at comparable viability when compared to needle injections15,16.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>50 mL centrifuge tube</td>
			<td>Greiner BioOne</td>
			<td>227261</td>
			<td></td>
		</tr>
		<tr>
			<td>1 mL BD Luer-LokTM Syringe</td>
			<td>BD Plastik Inc</td>
			<td>n.a.</td>
			<td></td>
		</tr>
		<tr>
			<td>100 &#181;m cell sieve</td>
			<td>Greiner BioOne</td>
			<td>542000</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL centrifuge tube</td>
			<td>Greiner BioOne</td>
			<td>188271</td>
			<td></td>
		</tr>
		<tr>
			<td>75 cm2 tissue culture flask</td>
			<td>Corning Incorporated</td>
			<td>353136</td>
			<td></td>
		</tr>
		<tr>
			<td>AFM head</td>
			<td>(CellHesion 200) JPK</td>
			<td>JPK00518</td>
			<td></td>
		</tr>
		<tr>
			<td>AFM processing software</td>
			<td>Bruker</td>
			<td>JPK00518</td>
			<td></td>
		</tr>
		<tr>
			<td>AFM software</td>
			<td>Bruker</td>
			<td>JPK00518</td>
			<td></td>
		</tr>
		<tr>
			<td>AFM system Cell Hesion 200</td>
			<td>Bruker</td>
			<td>JPK00518</td>
			<td></td>
		</tr>
		<tr>
			<td>All-In-One-Al cantilever</td>
			<td>Budget Sensors</td>
			<td>AIO-10</td>
			<td>tip A, Conatct Mode, Shape: Beam 
			Force Constant: 0.2 N/m (0.04 - 0.7 N/m) 
			Resonance Frequency: 15 kHz (10 - 20 kHz) 
			Length: 500 &#181;m (490 - 510 &#181;m) 
			Width: 30 &#181;m (35 - 45 &#181;m) 
			Thickness: 2.7 &#181;m (1.7 - 3.7 &#181;m)</td>
		</tr>
		<tr>
			<td>Amphotericin B solution</td>
			<td>Sigma</td>
			<td>A2942</td>
			<td>250 &#181;g/ml</td>
		</tr>
		<tr>
			<td>Atomic Force Microscope (AFM)</td>
			<td>CellHesion 200, JPK Instruments, Berlin, Germany</td>
			<td>JPK00518</td>
			<td></td>
		</tr>
		<tr>
			<td>BD Microlance 3 18G</td>
			<td>BD</td>
			<td>304622</td>
			<td></td>
		</tr>
		<tr>
			<td>bovine serum albumin</td>
			<td>Gibco</td>
			<td>A10008-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Cantilever</td>
			<td>&#160;All-In-One-AleTl, Budget Sensors, Sofia, Bulgaria</td>
			<td>AIO-TL-10</td>
			<td>tip A, k &#188; 0.2 N/m</td>
		</tr>
		<tr>
			<td>C-chip disposable hemocytometer</td>
			<td>NanoEnTek</td>
			<td>631-1098</td>
			<td></td>
		</tr>
		<tr>
			<td>centrifuge: Rotina 420R</td>
			<td>Hettich Zentrifugen</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase, Type I, powder</td>
			<td>Gibco</td>
			<td>17100-017</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s Modified Eagle&#8217;s Medium - low glucose</td>
			<td>Sigma</td>
			<td>D5546</td>
			<td></td>
		</tr>
		<tr>
			<td>Feather disposable scalpel (No. 10)</td>
			<td>Feather</td>
			<td>02.001.30.010</td>
			<td></td>
		</tr>
		<tr>
			<td>fetal bovine serum (FBS)</td>
			<td>Sigma</td>
			<td>F7524</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES sodium salt solution (1 M)</td>
			<td>Sigma</td>
			<td>H3662</td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted phase contrast microscope (Integrated with AFM)</td>
			<td>AxioObserver D1, Carl Zeiss Microscopy, Jena, Germany</td>
			<td>L201306_03</td>
			<td></td>
		</tr>
		<tr>
			<td>laboratory bags</td>
			<td>Brand</td>
			<td>759705</td>
			<td></td>
		</tr>
		<tr>
			<td>Leibovitz&#39;s L-15 medium without l-glutamine</td>
			<td>Merck</td>
			<td>F1315</td>
			<td></td>
		</tr>
		<tr>
			<td>Leibovitz&#39;s L-15 medium without L-glutamine</td>
			<td>(Merck KGaA, Darmstadt, Germany)</td>
			<td>F1315</td>
			<td></td>
		</tr>
		<tr>
			<td>L-glutamine</td>
			<td>Lonza</td>
			<td>BE 17-605C1</td>
			<td>200 mM</td>
		</tr>
		<tr>
			<td>LIVE/DEADTM Viability/Cytotoxicity Kit</td>
			<td>Invitrogen by Thermo Fisher Scientific</td>
			<td>L3224</td>
			<td>Calcein AM and EthD-1 are used from this kit.</td>
		</tr>
		<tr>
			<td>Microscope software: Zen 2.6</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope: AxioVertA.1</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Nelaton-Catheter female</td>
			<td>Bicakcilar</td>
			<td>19512051</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>15140-122</td>
			<td>10000 U/ml Penicillin 
			10000 &#181;g/ml Streptomycin</td>
		</tr>
		<tr>
			<td>Petri dish heater associated with AFM</td>
			<td>Bruker</td>
			<td>T-05-0117</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish heater associated with AFM</td>
			<td>JPK Instruments AG, Berlin, Germany</td>
			<td>T-05-0117</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS)</td>
			<td>Gibco</td>
			<td>10010-015</td>
			<td></td>
		</tr>
		<tr>
			<td>Statistical Software: SPSS Statistics 22</td>
			<td>IBM</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile Petri dish - CellStar</td>
			<td>Greiner BioOne</td>
			<td>664160</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue culture dishes</td>
			<td>TPP AG</td>
			<td>TPP93040</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue culture dishes</td>
			<td>TPP Techno Plastic Products AG, Trasadingen, Switzerland</td>
			<td>TPP93040</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan Blue 0.4% 
			0.85% NaCl</td>
			<td>Lonza</td>
			<td>17-942E</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin-EDTA solution</td>
			<td>Sigma</td>
			<td>T3924</td>
			<td></td>
		</tr>
		<tr>
			<td>Waterjet: ERBEJET2 device</td>
			<td>Erbe Elektromedizin GmbH</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Williams Cystoscopic Injection Needle</td>
			<td>Cook Medical</td>
			<td>G14220</td>
			<td>23G, 5.0 Fr, 35 cm</td>
		</tr>
	</tbody>
</table>

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%.

Following cell delivery via the two approaches, the viability of cells delivered through the WN (97.2 &#177; 2%, n=10, p&#60;0.002) was higher when compared to injections by WJ using the E60-10 settings (85.9 &#177; 0.16%, n=12) (Figure 2). Biomechanical assessment results showed that: WN injections of cells in capture fluid displayed no significant difference with respect to the elastic moduli (EM; 0.992 kPa) when compared to the controls (1.176 kPa; Figure 3A)...

Watch this Scientific Journal Video about Injection of Porcine Adipose Tissue-Derived Stroma Cells via Waterjet Technology at JoVE.com

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

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 ...

injection-porcine-adipose-tissue-derived-stroma-cells-via-waterjet

Research

JoVE Journal

Bioengineering

1.8K Views.  University Hospital Tübingen. 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.

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

Procedure for Decellularization of Porcine Heart by Retrograde Coronary Perfusion

Perfusion-based whole organ decellularization has recently gained interest in the field of tissue engineering as a means to create site-specific extracellular matrix scaffolds, while largely preserving the native architecture of the scaffold. To date, this approach has been utilized in a variety of organ systems, including the heart, lung, and liver 1-5. Previous decellularization methods for tissues without an easily accessible vascular network have relied upon prolonged exposure of tissue to solutions of detergents, acids, or enzymatic treatments as a means to remove the cellular and nuclear components from the surrounding extracellular environment6-8. However, the effectiveness of these methods hinged upon the ability of the solutions to permeate the tissue via diffusion. In contrast, perfusion of organs through the natural vascular system effectively reduced the diffusion distance and facilitated transport of decellularization agents into the tissue and cellular components out of the tissue. Herein, we describe a method to fully decellularize an intact porcine heart through coronary retrograde perfusion. The protocol yielded a fully decellularized cardiac extracellular matrix (c-ECM) scaffold with the three-dimensional structure of the heart intact. Our method used a series of enzymes, detergents, and acids coupled with hypertonic and hypotonic rinses to aid in the lysis and removal of cells. The protocol used a Trypsin solution to detach cells from the matrix followed by Triton X-100 and sodium deoxycholate solutions to aid in removal of cellular material. The described protocol also uses perfusion speeds of greater than 2 L/min for extended periods of time. The high flow rate, coupled with solution changes allowed transport of agents to the tissue without contamination of cellular debris and ensured effective rinsing of the tissue. The described method removed all nuclear material from native porcine cardiac tissue, creating a site-specific cardiac ECM scaffold that can be used for a variety of applications.

A method to rapidly and completely remove cellular components from an intact porcine heart through retrograde perfusion is described. This method yields a site specific cardiac extracellular matrix scaffold which has the potential for use in multiple clinical applications.

Perfusion-based  whole organ decellularization has recently gained interest in the field of  tissue engineering as a means to create site-specific ...

Transplantation of Induced Pluripotent Stem Cell-derived Mesoangioblast-like Myogenic Progenitors in Mouse Models of Muscle Regeneration

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells similar to pericyte-derived mesoangioblasts (iPSC-derived mesoangioblast-like stem/progenitor cells: IDEMs) can be established from iPSCs generated from patients affected by different forms of muscular dystrophy. Patient-specific IDEMs can be genetically corrected with different strategies (e.g. lentiviral vectors, human artificial chromosomes) and enhanced in their myogenic differentiation potential upon overexpression of the myogenesis regulator MyoD. This myogenic potential is then assessed in vitro with specific differentiation assays and analyzed by immunofluorescence. The regenerative potential of IDEMs is further evaluated in vivo, upon intramuscular and intra-arterial transplantation in two representative mouse models displaying acute and chronic muscle regeneration. The contribution of IDEMs to the host skeletal muscle is then confirmed by different functional tests in transplanted mice. In particular, the amelioration of the motor capacity of the animals is studied with treadmill tests. Cell engraftment and differentiation are then assessed by a number of histological and immunofluorescence assays on transplanted muscles. Overall, this paper describes the assays and tools currently utilized to evaluate the differentiation capacity of IDEMs, focusing on the transplantation methods and subsequent outcome measures to analyze the efficacy of cell transplantation.

Induced pluripotent stem cell (iPSC)-derived myogenic progenitors are promising candidates for cell therapy strategies to treat muscular dystrophies. This protocol describes transplantation and functional measurements required to evaluate the engraftment and differentiation of iPSC-derived mesoangioblasts (a type of muscle progenitors) in mouse models of acute and chronic muscle regeneration.

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells ...

Human Cartilage Tissue Fabrication Using Three-dimensional Inkjet Printing Technology

Bioprinting, which is based on thermal inkjet printing, is one of the most attractive enabling technologies in the field of tissue engineering and regenerative medicine. With digital control&#160;cells, scaffolds, and growth factors can be precisely deposited to the desired two-dimensional (2D) and three-dimensional (3D) locations rapidly. Therefore, this technology is an ideal approach to fabricate tissues mimicking their native anatomic structures. In order to engineer cartilage with native zonal organization, extracellular matrix composition (ECM), and mechanical properties, we developed a bioprinting platform using a commercial inkjet printer with simultaneous photopolymerization capable for 3D cartilage tissue engineering. Human chondrocytes suspended in poly(ethylene glycol) diacrylate (PEGDA) were printed for 3D neocartilage construction via layer-by-layer assembly. The printed cells were fixed at their original deposited positions, supported by the surrounding scaffold in simultaneous photopolymerization. The mechanical properties of the printed tissue were similar to the native cartilage. Compared to conventional tissue fabrication, which requires longer UV exposure, the viability of the printed cells with simultaneous photopolymerization was significantly higher. Printed neocartilage demonstrated excellent glycosaminoglycan (GAG) and collagen type II production, which was consistent with gene expression. Therefore, this platform is ideal for accurate cell distribution and arrangement for anatomic tissue engineering.

The methods described in this paper show how to convert a commercial inkjet printer into a bioprinter with simultaneous UV polymerization. The printer is capable of constructing 3D tissue structure with cells and biomaterials. The study demonstrated here constructed a 3D neocartilage.

Bioprinting, which is based on thermal inkjet printing, is one of the most attractive enabling technologies in the field of tissue engineering and ...

An Injectable and Drug-loaded Supramolecular Hydrogel for Local Catheter Injection into the Pig Heart

Regeneration of lost myocardium is an important goal for future therapies because of the increasing occurrence of chronic ischemic heart failure and the limited access to donor hearts. An example of a treatment to recover the function of the heart consists of the local delivery of drugs and bioactives from a hydrogel. In this paper a method is introduced to formulate and inject a drug-loaded hydrogel non-invasively and side-specific into the pig heart using a long, flexible catheter. The use of 3-D electromechanical mapping and injection via a catheter allows side-specific treatment of the myocardium. To provide a hydrogel compatible with this catheter, a supramolecular hydrogel is used because of the convenient switching from a gel to a solution state using environmental triggers. At basic pH this ureido-pyrimidinone modified poly(ethylene glycol) acts as a Newtonian fluid which can be easily injected, but at physiological pH the solution rapidly switches into a gel. These mild switching conditions allow for the incorporation of bioactive drugs and bioactive species, such as growth factors and exosomes as we present here in both in vitro and in vivo experiments. The in vitro experiments give an on forehand indication of the gel stability and drug release, which allows for tuning of the gel and release properties before the subsequent application in vivo. This combination allows for the optimal tuning of the gel to the used bioactive compounds and species, and the injection system.

Supramolecular hydrogelators based on ureido-pyrimidinones allow full control over the macroscopic gel properties and the sol&#8211;gel switching behavior using pH. Here, we present a protocol for formulating and injecting such a supramolecular hydrogelator via a catheter delivery system for local delivery directly in relevant areas in the pig heart.

Regeneration of lost myocardium is an important goal for future therapies because of the increasing occurrence of chronic ischemic heart failure and the ...

Isolation of Murine Adipose Tissue-derived Microvascular Fragments as Vascularization Units for Tissue Engineering

A functional microvascular network is of pivotal importance for the survival and integration of engineered tissue constructs. For this purpose, several angiogenic and prevascularization strategies have been established. However, most cell-based approaches include time-consuming in vitro steps for the formation of a microvascular network. Hence, they are not suitable for intraoperative one-step procedures. Adipose tissue-derived microvascular fragments (ad-MVF) represent promising vascularization units. They can be easily isolated from fat tissue and exhibit a functional microvessel morphology. Moreover, they rapidly reassemble into new microvascular networks after in vivo implantation. In addition, ad-MVF have been shown to induce lymphangiogenesis. Finally, they are a rich source of mesenchymal stem cells, which may further contribute to their high vascularization potential. In previous studies we have demonstrated the remarkable vascularization capacity of ad-MVF in engineered bone and skin substitutes. In the present study, we report on a standardized protocol for the enzymatic isolation of ad-MVF from murine fat tissue.

We present a protocol to isolate adipose tissue-derived microvascular fragments that represent promising vascularization units. They can be rapidly isolated, do not require in vitro processing and, thus, may be used for one-step prevascularization in different fields of tissue engineering.

A functional microvascular network is of pivotal importance for the survival and integration of engineered tissue constructs. For this purpose, several ...

Generic Protocol for Optimization of Heterologous Protein Production Using Automated Microbioreactor Technology

A core business in industrial biotechnology using microbial production cell factories is the iterative process of strain engineering and optimization of bioprocess conditions. One important aspect is the improvement of cultivation medium to provide an optimal environment for microbial formation of the product of interest. It is well accepted that the media composition can dramatically influence overall bioprocess performance. Nutrition medium optimization is known to improve recombinant protein production with microbial systems and thus, this is a rewarding step in bioprocess development. However, very often standard media recipes are taken from literature, since tailor-made design of the cultivation medium is a tedious task that demands microbioreactor technology for sufficient cultivation throughput, fast product analytics, as well as support by lab robotics to enable reliability in liquid handling steps. Furthermore, advanced mathematical methods are required for rationally analyzing measurement data and efficiently designing parallel experiments such as to achieve optimal information content.
The generic nature of the presented protocol allows for easy adaption to different lab equipment, other expression hosts, and target proteins of interest, as well as further bioprocess parameters. Moreover, other optimization objectives like protein production rate, specific yield, or product quality can be chosen to fit the scope of other optimization studies. The applied Kriging Toolbox (KriKit) is a general tool for Design of Experiments (DOE) that contributes to improved holistic bioprocess optimization. It also supports multi-objective optimization which can be important in optimizing both upstream and downstream processes.

This manuscript describes a generic approach for tailor-made design of microbial cultivation media. This is enabled by an iterative workflow combining Kriging-based experimental design and microbioreactor technology for sufficient cultivation throughput, which is supported by lab robotics to increase reliability and speed in liquid handling media preparation.

A core business in industrial biotechnology using microbial production cell factories is the iterative process of strain engineering and optimization of ...

Microdissection of Primary Renal Tissue Segments and Incorporation with Novel Scaffold-free Construct Technology

Kidney transplantation is now a mainstream therapy for end-stage renal disease. However, with approximately 96,000 people on the waiting list and only one-fourth of these patients achieving transplantation, there is a dire need for alternatives for those with failing organs. In order to decrease the harmful consequences of dialysis along with the overall healthcare costs it incurs, active investigation is ongoing in search of alternative solutions to organ transplantation. Implantable tissue-engineered renal cellular constructs are one such feasible approach to replacing lost renal functionality. Here, described for the first time, is the microdissection of murine kidneys for isolation of living corticomedullary renal segments. These segments are capable of rapid incorporation within scaffold-free endothelial-fibroblast constructs which may enable rapid connection with host vasculature once implanted. Adult mouse kidneys were procured from living donors, followed by stereoscope microdissection to obtain renal segments 200 - 300 &#181;m in diameter. Multiple renal constructs were fabricated using primary renal segments harvested from only one kidney. This method demonstrates a procedure which could salvage functional renal tissue from organs that would otherwise be discarded.

Tissue-engineered renal constructs provide a solution for the organ shortage and deleterious effects of dialysis. Here, we describe a protocol to micro dissect murine kidneys for isolation of cortico-medullary segments. These segments are implanted into scaffold-free cellular constructs, forming renal organoids.

Kidney transplantation is now a mainstream therapy for end-stage renal disease. However, with approximately 96,000 people on the waiting list and only ...

Isolation and Decellularization of a Whole Porcine Pancreas

Tissue engineering of the whole pancreas can improve current treatments for diabetes mellitus. The ultimate goal is to tissue engineer pancreas from an allogeneic or xenogeneic source with human cells. A demonstration of methods for the efficient dissection, decellularization, and recellularization of porcine pancreas might benefit the field. Akin to human pancreases, porcine pancreases have a special anatomical arrangement with three lobes (splenic, duodenal, and connection) rounded by the duodenum and small intestine. The duodenal lobe of the pancreas connects to the duodenum by several small blood vessels. Tissue engineering of the pancreas is complicated because of its exocrine and endocrine nature. In this paper, we show a detailed protocol to dissect the whole porcine pancreas and decellularize it with detergents while saving its structure and some extracellular matrix components. To achieve complete perfusion, the aorta is chosen as inlet and the portal vein as outlet. The other blood vessels (hepatic artery, splenic vein, splenic artery,&#160;mesenteric artery and vein tree) and bile duct are ligated. To prevent the formation of thrombus, the pig is heparinized and, immediately after dissection, the organ is flushed with cold heparin. To inhibit the action of exocrine enzymes, the pancreas decellularization is set at 4 &#176;C. The decellularization is performed by perfusion of Triton X-100, sodium deoxycholate, and deoxyribonuclease, with an intermittent and final extensive washing. With a successful decellularization, the pancreas appears white, and a histological evaluation with hematoxylin and eosin shows an absence of nuclei with a preserved extracellular matrix structure. Thus, the proposed method can be used to successfully dissect and decellularize whole porcine pancreas.

Tissue engineering of the whole pancreas is a challenge because of its exocrine and endocrine functions. We show a method for the dissection of an intact porcine pancreas and the process of successful decellularization by perfusion of detergents Triton X-100, sodium deoxycholate, and deoxyribonuclease.

Tissue engineering of the whole pancreas can improve current treatments for diabetes mellitus. The ultimate goal is to tissue engineer pancreas from an ...

High-resolution Imaging of Nuclear Dynamics in Live Cells under Uniaxial Tensile Strain

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the effect of applied extracellular tensile strain on cell populations in vitro via biochemical assays, a device has previously been designed which can be fabricated simply and is small enough to fit inside tissue culture incubators, as well as on top of microscope stages. However, the previous design of the polydimethylsiloxane substratum did not allow high-resolution subcellular imaging via oil-immersion objectives. This work describes a redesigned geometry of the polydimethylsiloxane substratum and a customized imaging setup that together can facilitate high-resolution subcellular imaging of live cells while under applied strain. This substratum can be used with the same, earlier designed device and, hence, has the same advantages as listed above, in addition to allowing high-resolution optical imaging. The design of the polydimethylsiloxane substratum can be improved by incorporating a grid which will facilitate tracking the same cell before and after the application of strain. Representative results demonstrate high-resolution time-lapse imaging of fluorescently labeled nuclei within strained cells captured using the method described here. These nuclear dynamics data give insights into the mechanism by which applied tensile strain promotes differentiation of oligodendrocyte progenitor cells.&#160;

Using a previously designed device to apply mechanical strain to adherent cells, this paper describes a redesigned substratum geometry and a customized apparatus for high-resolution single-cell imaging of strained cells with a 100x oil immersion objective.

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the ...

Pancreatic Tissue-Derived Extracellular Matrix Bioink for Printing 3D Cell-Laden Pancreatic Tissue Constructs

The transplantation of pancreatic islets is a promising treatment for patients who suffer from type 1 diabetes accompanied by hypoglycemia and secondary complications. However, islet transplantation still has several limitations such as the low viability of transplanted islets due to poor islet engraftment and hostile environments. In addition, the insulin-producing cells differentiated from human pluripotent stem cells have limited ability to secrete sufficient hormones that can regulate the blood glucose level; therefore, improving the maturation by culturing cells with proper microenvironmental cues is strongly required. In this article, we elucidate protocols for preparing a pancreatic tissue-derived decellularized extracellular matrix (pdECM) bioink to provide a beneficial microenvironment that can increase glucose sensitivity of pancreatic islets, followed by describing the processes for generating 3D pancreatic tissue constructs using a microextrusion-based bioprinting technique.

Decellularized extracellular matrix (dECM) can provide suitable microenvironmental cues to recapitulate the inherent functions of target tissues in an engineered construct. This article elucidates the protocols for the decellularization of pancreatic tissue, evaluation of pancreatic tissue-derived dECM bioink, and generation of 3D pancreatic tissue constructs using a bioprinting technique.

The transplantation of pancreatic islets is a promising treatment for patients who suffer from type 1 diabetes accompanied by hypoglycemia and secondary ...