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: more than 80% viable cells post delivery18. While a standardized and reproducible protocol is achieved with the WJ approach, the outcome of needle injection delivery is highly dependent on the size and nozzle of the syringe and needle, pressure, flow rate and the physician performing the injection themselves19.
Studies employing WJ cell delivery in living animal models showed that by varying ejection pressure, the penetration depth can be adapted to the targeted tissue and as such, to the desired clinical application13,16. Transurethral cell injections in living animals under visual control reported misplacement or loss of cells in about 50% of animals treated20, while WJ injections reported precise cell injection rates above 90% (Linzenbold et al.16 and unpublished observation). The current golden standard for cell delivery (needle injections) require penetration of the cannula in a targeted tissue. Therefore, needle cell translation causes injury and trauma in all cases. In the urethra, this may actually cause inflammation ad toxification due to germ and toxins found in even in healthy urine. Additionally, the user-operation time in WJ injection is significantly shorter when compared to a needle injection: cells are placed within milliseconds into the intended tissue layer by presetting the pressure levels. In contrast, penetration depth on needle injection in remote areas by endoscopy is dependent on the skills and experience of the surgeon. The reproducibility of WJ injection is also expected to be superior, but presently only pre-clinical data exist15,16,20 and less than 200 animals were investigated. In our recent study we noted that cellular elasticity is reduced by WJ application when compared to needle injections21. This can be attributed to the shear stress of cells in higher velocity during WJ delivery. Moreover, eventual cell loss might be compensated by a higher precision of cell placement and ejection within the region of interest as achieved with WJ guided delivery by visual guided cystoscopy22.
It is well known that mechanical forces direct stem cell behavior, regeneration potential as well as their subsequent viability and functionality post-transplantation 10,23. The advent of atomic AFM provided a powerful tool for quantifying the mechanical properties of single living cells in nano scale resolution in aqueous conditions 24,25,26,27. AFM is a reliable and highly sensitive method that can detect and record stiffnesses ranging from less than 100 Pa to 106 Pa, thus covering a wide range for the majority tissues and cells28. In fact, cell mechanics is emerging as label-free biomarker for evaluating cell state and in both physiological and pathological state29 and cell elasticity is the synergetic and cumulative response of the nucleus - cytoskeleton crosstalk. It is well established that when a cell is subjected to external forces, these forces are transmitted from the plasma membrane via the cytoskeleton to the nucleus, resulting in intra-nuclear deformations and reorganization30,31,32. Therefore the nucleus, long seen as the genomic material and transcription apparatus, is a key player in the cellular mechanotransduction as well32. In fact, the importance of nuclear mechanics and nucleo-cytoskeletal connections, in all cellular functions and mutations, in lamins and linkers of the nucleo-skeleton to the cytoskeleton (LINC) complex - are at the onset foundation of several pathologies 33,34. These forces could also indicate cellular artifacts. Specifically, external generated forces actually propagate along cytoskeletal filaments and are further transmitted to the nuclear lamina across the LINC complex; in response to these forces, the nucleus, in fact, becomes stiffer35,36, thus merging the two closely intertwined and connected processes. This is also the rationale of our approach and our nuclear mechanics measurements. Moreover, a precise placement of the cantilever on top of the nuclear surface also ensures a high degree of reproducibility and reduces variations owing to cell heterogeneity and attachment to the substratum.
Even though cell elasticity is emerging as label-free biomarker for evaluating the cell state and in both physiological and pathological states29, the measured elastic moduli are marked by large variations even in the same cell type37. A method to counteract such variations as suggested by Schillers et al. is the implementation of standardized nanomechanical AFM procedures (SNAP) that ensure a high reproducibility and applicability of elasticity measurements as a reliable quantitative marker to cells in various states38. Also, even though experimental parameters employed in the AFM analyses, such as indentation velocity, indenter shape and size as well as accurate representation of tip geometry in model fitting 39, influence the absolute measured values40,41, these parameters should not impact the results within one study or a measured tendency.
However, bear in mind that AFM indentations are restricted to the analysis of the outer surface of cells and are, thus, incapable of scanning the inside of a cell membrane or particular intracellular structures. Usukura et al. proposed a &#34;unroofing&#34; method that breaks the cellular membrane and removes the cytoplasmic-soluble components42, thus allowing AFM- intracellular investigations. In our study, however, the focus was placed on the assessment of average elastic moduli rather than probing distinct and selective intracellular components.
Collectively, the consistency and reliability of the yielded AFM data strongly depends on the technical experience of the respective operator and could be biased by biological variability38. Accounting for all the sensitive variables that might affect the actual AFM results, the absolute elastic values reported in this study cannot be generalized and are rather specific for our experimental setup.
Overall, our study provides evidence21 as well as a step-by-step protocol for the superiority of WJ injections over needle injections for regenerative cell therapy regimen.

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><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 &amp;micro;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 cm&lt;sup&gt;2&lt;/sup&gt; 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&lt;br/&gt; Force Constant: 0.2 N/m (0.04 - 0.7 N/m)&lt;br/&gt; Resonance Frequency: 15 kHz (10 - 20 kHz)&lt;br/&gt; Length: 500 &amp;micro;m (490 - 510 &amp;micro;m)&lt;br/&gt; Width: 30 &amp;micro;m (35 - 45 &amp;micro;m)&lt;br/&gt; Thickness: 2.7 &amp;micro;m (1.7 - 3.7 &amp;micro;m)</td></tr><tr><td>Amphotericin B solution</td><td>Sigma</td><td>A2942</td><td>250 &amp;micro;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>&amp;nbsp;All-In-One-AleTl, Budget Sensors, Sofia, Bulgaria</td><td>AIO-TL-10</td><td>tip A, k &amp;frac14; 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&amp;rsquo;s Modified Eagle&amp;rsquo;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&#039;s L-15 medium without l-glutamine</td><td>Merck</td><td>F1315</td><td></td></tr><tr><td>Leibovitz&#039;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&lt;br/&gt; 10000 &amp;micro;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%&lt;br/&gt; 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), while WJ injections triggered a significant reduction of the cellular EM (0.440 kPa, p&#60;0.001, Figure 3B). A decrease of 40 - 50% of the EM after WJ injections was noted. Even though, WN injections in cadaveric urethra tissue yielded no significant difference in cellular EM (Figure 4A) a significant reduction in EM was noted after WJ injections in tissue samples (0.890 kPa to 0.429 kPa; p&#60;0.00, Figure 4B). Thus, absolute EM values after WJ injection were thereby reduced by 51%. Collectively, the results show that while WJ cell delivery fulfills an absolute requirement for a clinical implementation where more than 80% viable cells post delivery18 , post WJ delivery the cell elastic moduli are affected. A lower cellular EM might facilitate the migration of features of the cells after WJ delivery. In such a wider distribution and range of regenerative capacities in the desired region19.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63132/63132fig01.jpg" /> 
Figure 1. Anatomy of porcine bladder and urethra and injection sites. A) The ventral side of the porcine bladder and urethra with the three ligaments fixing the bladder in the abdominal and pelvic cavity are shown. Additionally, the ureters that end on the dorsal side of the bladder are shown. B) Representative image of the cadaveric urethra used for WJ and WN injection. Longitudinally, the dorsal opened urethra with injection domes circled in black are shown. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63132/63132fig01large.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/63132/63132fig02.jpg" /> 
Figure 2. Cellular viability determination injections via WN and WJ. pADSCs injected via WN or WJ were collected after injection and counted via Trypan exclusion to determine the viability. The cellular viability was significantly reduced after WJ injection when compared to cells delivered via WN. *** p&#60;0.001. The data is graphically displayed as mean with standard deviation. Abbreviations: WJ - waterjet, WN - Williams needle. Figure adapted from Danalache et al. 202119. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63132/63132fig02large.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/63132/63132fig03.jpg" /> 
Figure 3. Comparison of the quantified Young&#39;s moduli of WN respectively WJ delivered cells into capture media and their corresponding controls. No notable difference in EM was observed in the boxplots for the control (untreated) cell monolayers and WN delivered cells (A). Contrastingly, a significant decrease in elasticity can be noted between the boxplots of the control cells and the WJ group (B). ns - not significant, p &#62; 0.05, ***p&#60;0.001. Abbreviations: WJ - waterjet, WN - Williams needle. Figure adapted from Danalache et al. 202119. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63132/63132fig03large.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/63132/63132fig04.jpg" /> 
Figure 4. Comparison of the quantified Young&#39;s moduli of WN respectively WJ delivered cells into cadaveric urethra and their corresponding controls. No notable difference was observed between cells delivered via WN cells and their corresponding controls (A). A significant decrease in elasticity was noted between the WJ delivered cells and the control cell monolayer (B). ns - not significant, p &#62; 0.05, ***p&#60;0.001. Abbreviations: WJ - waterjet, WN - Williams needle. Figure adapted from Danalache et al. 202119. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63132/63132fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Approach parameter</td>
			<td>Value</td>
		</tr>
		<tr>
			<td>Approach IGain</td>
			<td>3.0 Hz</td>
		</tr>
		<tr>
			<td>Approach PGain</td>
			<td>0.0002</td>
		</tr>
		<tr>
			<td>Approach target height</td>
			<td>10.0 &#181;m</td>
		</tr>
		<tr>
			<td>Approach setpoint</td>
			<td>3.00 V</td>
		</tr>
		<tr>
			<td>Approach baseline</td>
			<td>0.00 V</td>
		</tr>
	</tbody>
</table>
Table 1. Approach parameters.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Run parameter</td>
			<td>Value</td>
		</tr>
		<tr>
			<td>Set point</td>
			<td>10 nN</td>
		</tr>
		<tr>
			<td rowspan="2">Z Movement/ Extend Speed</td>
			<td>Constant speed/</td>
		</tr>
		<tr>
			<td>5.0 &#181;m/s</td>
		</tr>
		<tr>
			<td>Contact time</td>
			<td>0.0 s</td>
		</tr>
		<tr>
			<td>Pulling length</td>
			<td>90 &#181;m</td>
		</tr>
		<tr>
			<td>Delay Mode</td>
			<td>Constant Force</td>
		</tr>
		<tr>
			<td>Sample rate</td>
			<td>2000 Hz</td>
		</tr>
	</tbody>
</table>
Table 2. Run parameters.

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

Nanyang Technological University

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

Isolation and Enrichment of Human Adipose-derived Stromal Cells for Enhanced Osteogenesis

Bone marrow-derived mesenchymal stromal cells (BM-MSCs) are considered the gold standard for stem cell-based tissue engineering applications. However, the process by which they must be harvested can be associated with significant donor site morbidity. In contrast, adipose-derived stromal cells (ASCs) are more readily abundant and more easily harvested, making them an appealing alternative to BM-MSCs. Like BM-MSCs, ASCs can differentiate into osteogenic lineage cells and can be used in tissue engineering applications, such as seeding onto scaffolds for use in craniofacial skeletal defects. ASCs are obtained from the stromal vascular fraction (SVF) of digested adipose tissue, which is a heterogeneous mixture of ASCs, vascular endothelial and mural cells, smooth muscle cells, pericytes, fibroblasts, and circulating cells. Flow cytometric analysis has shown that the surface marker profile for ASCs is similar to that for BM-MSCs. Despite several published reports establishing markers for the ASC phenotype, there is still a lack of consensus over profiles identifying osteoprogenitor cells in this heterogeneous population. This protocol describes how to isolate and use a subpopulation of ASCs with enhanced osteogenic capacity to repair critical-sized calvarial defects.

The transcriptional heterogeneity within human adipose-derived stromal cells can be defined on the single cell level using cell surface markers and osteogenic genes. We describe a protocol utilizing flow cytometry for the isolation of cell subpopulations with increased osteogenic potential, which may be used to enhance craniofacial skeletal reconstruction.

Bone marrow-derived mesenchymal stromal cells (BM-MSCs) are considered the gold standard for stem cell-based tissue engineering applications. However, the ...

Developmental Biology

Use of Human Perivascular Stem Cells for Bone Regeneration

Human perivascular stem cells (PSCs) can be isolated in sufficient numbers from multiple tissues for purposes of skeletal tissue engineering1-3. PSCs are a FACS-sorted population of 'pericytes' (CD146+CD34-CD45-) and 'adventitial cells' (CD146-CD34+CD45-), each of which we have previously reported to have properties of mesenchymal stem cells. PSCs, like MSCs, are able to undergo osteogenic differentiation, as well as secrete pro-osteogenic cytokines1,2. In the present protocol, we demonstrate the osteogenicity of PSCs in several animal models including a muscle pouch implantation in SCID (severe combined immunodeficient) mice, a SCID mouse calvarial defect and a femoral segmental defect (FSD) in athymic rats. The thigh muscle pouch model is used to assess ectopic bone formation. Calvarial defects are centered on the parietal bone and are standardly 4 mm in diameter (critically sized)8. FSDs are bicortical and are stabilized with a polyethylene bar and K-wires4. The FSD described is also a critical size defect, which does not significantly heal on its own4. In contrast, if stem cells or growth factors are added to the defect site, significant bone regeneration can be appreciated. The overall goal of PSC xenografting is to demonstrate the osteogenic capability of this cell type in both ectopic and orthotopic bone regeneration models.

Human perivascular stem cells (PSCs) are a novel stem cell class for skeletal tissue regeneration similar to mesenchymal stem cells (MSCs). PSCs can be isolated by FACS (fluorescence activated cell sorting) from adipose tissue procured during standard liposuction procedures, then combined with an osteoinductive scaffold to achieve bone formation in vivo.

Human perivascular stem cells (PSCs) can be isolated in sufficient numbers from multiple tissues for purposes of skeletal tissue engineering1-3. PSCs are ...

Isolation and Differentiation of Adipose-Derived Stem Cells from Porcine Subcutaneous Adipose Tissues

Obesity is an unconstrained worldwide epidemic. Unraveling molecular controls in adipose tissue development holds promise to treat obesity or diabetes. Although numerous immortalized adipogenic cell lines have been established, adipose-derived stem cells from the stromal vascular fraction of subcutaneous white adipose tissues provide a reliable cellular system ex vivo much closer to adipose development in vivo. Pig adipose-derived stem cells (pADSC) are isolated from 7- to 9-day old piglets. The dorsal white fat depot of porcine subcutaneous adipose tissues is sliced, minced and collagenase digested. These pADSC exhibit strong potential to differentiate into adipocytes. Moreover, the pADSC also possess multipotency, assessed by selective stem cell markers, to differentiate into various mesenchymal cell types including adipocytes, osteocytes, and chondrocytes. These pADSC can be used for clarification of molecular switches in regulating classical adipocyte differentiation or in direction to other mesenchymal cell types of mesodermal origin. Furthermore, extended lineages into cells of ectodermal and endodermal origin have recently been achieved. Therefore, pADSC derived in this protocol provide an abundant and assessable source of adult mesenchymal stem cells with full multipotency for studying adipose development and application to tissue engineering of regenerative medicine.

This protocol describes the isolation of pig adipose-derived stem cells (pADSC) from subcutaneous adipose tissues with examination of multipotency. The multipotent pADSC are used to delineate processes of adipocyte differentiation and study transdifferentiation into multiple cell lineages of mesodermal mesenchyme or further lineages of ectoderm and endoderm for regenerative studies.

Obesity is an unconstrained worldwide epidemic. Unraveling molecular controls in adipose tissue development holds promise to treat obesity or diabetes. ...

Repair of a Critical-sized Calvarial Defect Model Using Adipose-derived Stromal Cells Harvested from Lipoaspirate

Craniofacial skeletal repair and regeneration offers the promise of de novo tissue formation through a cell-based approach utilizing stem cells. Adipose-derived stromal cells (ASCs) have proven to be an abundant source of multipotent stem cells capable of undergoing osteogenic, chondrogenic, adipogenic, and myogenic differentiation. Many studies have explored the osteogenic potential of these cells in vivo with the use of various scaffolding biomaterials for cellular delivery. It has been demonstrated that by utilizing an osteoconductive, hydroxyapatite-coated poly(lactic-co-glycolic acid) (HA-PLGA) scaffold seeded with ASCs, a critical-sized calvarial defect, a defect that is defined by its inability to undergo spontaneous healing over the lifetime of the animal, can be effectively show robust osseous regeneration. This in vivo model demonstrates the basis of translational approaches aimed to regenerate the bone tissue - the cellular component and biological matrix. This method serves as a model for the ultimate clinical application of a progenitor cell towards the repair of a specific tissue defect.

This protocol describes the isolation of adipose-derived stromal cells from lipoaspirate and the creation of a 4 mm critical-sized calvarial defect to evaluate skeletal regeneration.

Craniofacial skeletal repair and regeneration offers the promise of de novo tissue formation through a cell-based approach utilizing stem cells. ...

Medicine

A Microphysiologic Platform for Human Fat: Sandwiched White Adipose Tissue

White adipose tissue (WAT) plays a crucial role in regulating weight and everyday health. Still, there are significant limitations to available primary culture models, all of which have failed to faithfully recapitulate the adipose microenvironment or extend WAT viability beyond two weeks. The lack of a reliable primary culture model severely impedes research in WAT metabolism and drug development. To this end we have utilized NIH's standards of a microphysiologic system to develop a novel platform for WAT primary culture called 'SWAT' (sandwiched white adipose tissue). We overcome the natural buoyancy of adipocytes by sandwiching minced WAT clusters between sheets of adipose-derived stromal cells. In this construct, WAT samples are viable over eight weeks in culture. SWAT maintains the intact ECM, cell-to-cell contacts, and physical pressures of in vivo WAT conditions; additionally, SWAT maintains a robust transcriptional profile, sensitivity to exogenous chemical signaling, and whole tissue function. SWAT represents a simple, reproducible, and effective method of primary adipose culture. Potentially, it is a broadly applicable platform for research in WAT physiology, pathophysiology, metabolism, and pharmaceutical development.

White adipose tissue (WAT) has critical deficiencies in its current primary culture models, hindering pharmacological development and metabolic studies. Here, we present a protocol to produce an adipose microphysiological system by sandwiching WAT between sheets of stromal cells. This construct provides a stable and adaptable platform for primary WAT culture.

White adipose tissue (WAT) plays a crucial role in regulating weight and everyday health.  Still, there are significant limitations to available primary ...

Behavior

Creation and Transplantation of an Adipose-derived Stem Cell (ASC) Sheet in a Diabetic Wound-healing Model

Artificial skin has achieved considerable therapeutic results in clinical practice. However, artificial skin treatments for wounds in diabetic patients with impeded blood flow or with large wounds might be prolonged. Cell-based therapies have appeared as a new technique for the treatment of diabetic ulcers, and cell-sheet engineering has improved the efficacy of cell transplantation. A number of reports have suggested that adipose-derived stem cells (ASCs), a type of mesenchymal stromal cell (MSC), exhibit therapeutic potential due to their relative abundance in adipose tissue and their accessibility for collection when compared to MSCs from other tissues. Therefore, ASCs appear to be a good source of stem cells for therapeutic use. In this study, ASC sheets from the epididymal adipose fat of normal Lewis rats were successfully created using temperature-responsive culture dishes and normal culture medium containing ascorbic acid. The ASC sheets were transplanted into Zucker diabetic fatty (ZDF) rats, a rat model of type 2 diabetes and obesity, that exhibit diminished wound healing. A wound was created on the posterior cranial surface, ASC sheets were transplanted into the wound, and a bilayer artificial skin was used to cover the sheets. ZDF rats that received ASC sheets had better wound healing than ZDF rats without the transplantation of ASC sheets. This approach was limited because ASC sheets are sensitive to dry conditions, requiring the maintenance of a moist wound environment. Therefore, artificial skin was used to cover the ASC sheet to prevent drying. The allogenic transplantation of ASC sheets in combination with artificial skin might also be applicable to other intractable ulcers or burns, such as those observed with peripheral arterial disease and collagen disease, and might be administered to patients who are undernourished or are using steroids. Thus, this treatment might be the first step towards improving the therapeutic options for diabetic wound healing.

Creation and Transplantation of an Adipose-derived Stem Cell ASC Sheet in a Diabetic Wound-healing Model

Adipose-derived stem cells (ASCs) are easily isolated and harvested from the fat of normal rats. ASC sheets can be created using cell-sheet engineering and can be transplanted into Zucker diabetic fatty rats exhibiting full-thickness skin defects with exposed bone and then covered with a bilayer of artificial skin.

Artificial skin has achieved considerable therapeutic results in clinical practice. However, artificial skin treatments for wounds in diabetic patients ...

Clinical Protocol of Producing Adipose Tissue-Derived Stromal Vascular Fraction for Potential Cartilage Regeneration

Osteoarthritis (OA) is one of the most common debilitating disorders.&#160;Recently, numerous attempts have been made to improve the functions of the knees by using different forms of mesenchymal stem cells (MSCs). In Korea, bone marrow concentrates and cord blood-derived stem cells have been approved by the Korean Food and Drug Administration (KFDA) for cartilage regeneration. In addition, an adipose tissue-derived stromal vascular fraction (SVF) has been allowed by the KFDA for joint injections in human patients. Autologous adipose tissue-derived SVF contains extracellular matrix (ECM) in addition to mesenchymal stem cells. ECM excretes various cytokines that, along with hyaluronic acid (HA) and platelet-rich plasma (PRP) activated by calcium chloride, may help MSCs to regenerate cartilage and improve knee functions. In this article, we presented a protocol to improve knee functions by regenerating cartilage-like tissue in human patients with OA. The result of the protocol was first reported in 2011 followed by a few additional publications. The protocol involves liposuction to obtain autologous lipoaspirates that are mixed with collagenase. This lipoaspirates-collagenase mixture is then cut and homogenized to remove large fibrous tissue that may clog up the needle during the injection. Afterwards, the mixture is incubated to obtain adipose tissue-derived SVF. The resulting adipose tissue-derived SVF, containing both adipose tissue-derived MSCs and remnants of ECM, is injected into knees of patients, combined with HA and calcium chloride activated PRP. Included are three cases of patients who were treated with our protocol resulting in improvement of knee pain, swelling, and range of motion along with MRI evidence of hyaline cartilage-like tissue.

Here, we present a protocol to produce an adipose tissue-derived stromal vascular fraction and its application to improve knee functions by regenerating cartilage-like tissue in human patients with osteoarthritis.

Osteoarthritis (OA) is one of the most common debilitating disorders.&#160;Recently, numerous attempts have been made to improve the functions of the ...

Mechanical Micronization of Lipoaspirates for Regenerative Therapy

Stromal vascular fraction (SVF) has become a regenerative tool for various diseases; however, legislation strictly regulates the clinical application of cell products using collagenase. Here, we present a protocol to generate an injectable mixture of SVF cells and native extracellular matrix from adipose tissue by a purely mechanical process. Lipoaspirates are put into a centrifuge and spun at 1,200 x g for 3 min. The middle layer is collected and separated into two layers (high-density fat at the bottom and low-density fat on the top). The upper layer is directly emulsified by intersyringe shifting, at a rate of 20 mL/s for 6x to 8x. The emulsified fat is centrifuged at 2,000 x g for 3 min, and the sticky substance under the oil layer is collected and defined as the extracellular matrix (ECM)/SVF-gel. The oil on the top layer is collected. Approximately 5 mL of oil is added to 15 mL of high-density fat and emulsified by intersyringe shifting, at a rate of 20 mL/s for 6x to 8x. The emulsified fat is centrifuged at 2,000 x g for 3 min, and the sticky substance is also ECM/SVF-gel. After the transplantation of the ECM/SVF-gel into nude mice, the graft is harvested and assessed by histologic examination. The result shows that this product has the potential to regenerate into normal adipose tissue. This procedure is a simple, effective mechanical dissociation procedure to condense the SVF cells embedded in their natural supportive ECM for regenerative purposes.

Here, we present a protocol to obtain stromal vascular fraction from adipose tissue through a series of mechanical processes, which include emulsification and multiple centrifugations.

Stromal vascular fraction (SVF) has become a regenerative tool for various diseases; however, legislation strictly regulates the clinical application of ...

Technique for Obtaining Mesenchymal Stem Cell from Adipose Tissue and Stromal Vascular Fraction Characterization in Long-Term Cryopreservation

Human mesenchymal stem cells derived from adipose tissue have become increasingly attractive as they show appropriate features and are an accessible source for regenerative clinical applications. Different protocols have been used to obtain adipose-derived stem cells. This article describes different steps of an improved time-saving protocol to obtain a more significant amount of ADSC, showing how to cryopreserve and thaw ADSC to obtain viable cells for culture expansion. One hundred milliliters of lipoaspirate were collected, using a 26 cm three-hole and 3 mm caliber syringe liposuction, from the abdominal area of nine patients who subsequently underwent elective abdominoplasty. The stem cells isolation was carried out with a series of washes with Dulbecco&#39;s Phosphate Buffered Saline (DPBS) solution supplemented with calcium and the use of collagenase. Stromal Vascular Fraction (SVF) cells were cryopreserved, and their viability was checked by immunophenotyping. The SVF cellular yield was 15.7 x 105 cells/mL, ranging between 6.1-26.2 cells/mL. Adherent SVF cells reached confluence after an average of 7.5 (&#177;4.5) days, with an average cellular yield of 12.3 (&#177; 5.7) x 105 cells/mL. The viability of thawed SVF after 8 months, 1 year, and 2 years ranged between 23.06%-72.34% with an average of 47.7% (&#177;24.64) with the lowest viability correlating with cases of two-year freezing. The use of DPBS solution supplemented with calcium and bag resting times for fat precipitation with a shorter time of collagenase digestion resulted in an increased stem cell final cellular yield. The detailed procedure for obtaining high yields of viable stem cells was more efficient regarding time and cellular yield than the techniques from previous studies. Even after a long period of cryopreservation, viable ADSC cells were found in the SVF.

The present protocol describes an improved methodology for ADSC isolation resulting in a tremendous cellular yield with time gain compared to the literature. This study also provides a straightforward method for obtaining a relatively large number of viable cells after long-term cryopreservation.

Human mesenchymal stem cells derived from adipose tissue have become increasingly attractive as they show appropriate features and are an accessible ...

Mechanical Stromal Vascular Fraction in Fibrin Hydrogel for Regenerative Medicine

The Combination of Mechanically Isolated Stromal Vascular Fraction and Fibrin Hydrogel: A Processing Protocol

The regenerative potential of adipose-derived stromal cells (ASCs) has gained significant attention in regenerative and translational research. In the past, the extraction of these cells from adipose tissue required a multistep enzyme-based process, resulting in a heterogenous cell mix consisting of ACSs and other cells, which are jointly termed the stromal vascular fraction (SVF). More recently introduced mechanical SVF (mSVF) isolation protocols are less time-consuming and bypass regulatory concerns. We recently proposed a protocol that generates mSVF rich in stromal cells based on a combination of emulsification and centrifugation. One current issue in mSVF application for wound therapy application is the lack of a scaffold providing protection from mechanical manipulation and desiccation. Fibrin hydrogels have been shown to be a useful adjunct in cell transfer for wound healing purposes in the past. In the work herein, we delineate the preparation steps of an mSVF-fibrin hydrogel construct as a novel approach for translational research and clinical application.

Author Spotlight: Exploring the Potential of Fat-Derived Stromal Vascular Fraction for Wound Healing

Mechanically isolated stromal vascular fraction (SVF) in combination with a fibrin hydrogel offers an easy and efficient carrier for viable adipose-derived stromal cells for various indications, including tissue engineering and or wound healing purposes. Here, we present the preparation of a mechanical SVF (mSVF)-fibrin hydrogel construct for translational research and clinical application.

The regenerative potential of adipose-derived stromal cells (ASCs) has gained significant attention in regenerative and translational research. In the ...