The research leading to these results has received funding from the European Union Seventh Framework Programme FP7/2007-2013 under grant agreement n&#176; 607051 (BIO-INSPIRE). We thank Carolien van Spreuwel-Goossens from Fujifilm Manufacturing Europe B.V., for the technical assistance during RCP manufacturing, and Werner Stracke from Fraunhofer Institute for Silicate Research ISC, for assistance with the SEM analysis.

hBMSCs were isolated from the femur head of osteoarthritis patients undergoing femur head replacement surgery. The procedure was performed under the approval of the Local Ethics Committee of the University of Wuerzburg and informed consent of the patients. Primary microvascular endothelial cells were isolated from foreskin biopsies of juvenile donors. Their legal representative(s) provided full informed consent in writing. The study was approved by the local ethical board of the University of Wuerzburg (vote 182/10).
1. Isolation of hBMSCs and HDMECs
<ol>
	<li>Human bone marrow-derived mesenchymal stromal cells (hBMSCs)
	<ol>
		<li>Remove spongy bone from the femur head by using a sterile spoon. 
		NOTE: The spongy bone appears as a loose and granular substance inside the cortical bone, which is hard and rigid. It may be necessary to scratch parts of the spongy bone from the cortical bone by using a scalpel.</li>
		<li>Insert the removed spongy bone in two 50 mL tubes. The volume of the spongy bone can vary from 5 to 10 mL according to the dimension of the removed femur head during the hip replacement surgery.</li>
		<li>Wash with DMEM-F12 (1:1 mixture of DMEM and Ham F12 medium), filling the tube with approx. 30 mL of medium.</li>
		<li>Shake the tube manually, in a longitudinal way, for 5 s to let adipose cells out of the spongy bone. Centrifuge at 300 x g and 22 &#176;C for 5 min.</li>
		<li>Aspirate with a graduated plastic pipette, the fat-cell layer and approx. 20 mL of the remaining supernatant. Let enough medium to cover the spongy bone.</li>
		<li>Fill the tubes with 10 ml of DMEM-F12 and shake vigorously longitudinally by hand, for 10 - 15 s, to let the cells out.</li>
		<li>Transfer 10 - 15 mL of cell suspension with a graduated plastic pipette from the tubes to a new 50 mL tube.</li>
		<li>Repeat 2 - 3 times from step 1.1.6 pooling together the obtained cell suspension until the spongy bone collected in the 50 mL tube is white.</li>
		<li>Transfer the cell suspension with a graduated plastic pipette to two new tubes, avoiding aspirating the collagen pellet deposited on the bottom of the 50 mL tubes.</li>
		<li>Centrifuge the cell suspension at 300 x g and 22 &#176;C for 5 min. Discard the supernatant by aspirating it with a graduated plastic pipette.</li>
		<li>Add 40 mL of DMEM-F12 10% FCS 1% Pen/Strep 50 &#181;g/mL Ascorbate-2-phosphate in a third tube. Transfer 5 mL of this medium to each tube containing the cell pellet. Re-suspend the cell pellets by flickering the bottom of the tubes vigorously and transfer the cell suspensions to the tube containing only medium.</li>
		<li>OPTIONAL: Dilute 100 &#181;L of cells suspension for cell counting, diluting first 1:100 with PBS and then 1:10 in trypan blue, by adding 15 &#181;L of the first dilution to 15 &#181;L of DPBS and 30 &#181;L Trypan Blue to obtain a DPBS: Trypan Blue ratio of 1:1. Count the cells with a standard Neubauer chamber.</li>
		<li>Seed the cells at 109 cells/175 cm2 T-Flask in 25 mL of medium as described in 1.1.11. Alternatively, divide the whole cell suspension in up to ten 175 cm2 cell culture flasks without counting. At this stage, hBMSCs are not distinguishable from the erythrocytes, which greatly outnumber them. hBMSCs will appear as sparse colonies (1 - 2/flask) over the course of the first 7 days.</li>
		<li>Change medium after 4 days from seeding to allow interaction of the blood cells with the hBMSCs. After the first medium exchange, change the medium every 2 - 3 days. Perform the medium exchange by completely removing the culture medium with a graduated plastic pipette, washing two times with DPBS (by adding 10 mL of DPBS to the adherent cells and removing completely the DPBS with a graduated plastic pipette) and then adding 20 mL of fresh medium, as described in 1.1.11.</li>
	</ol>
	</li>
	<li>Human dermal microvascular endothelial cells (HDMECs)
	<ol>
		<li>Treat the skin biopsies and isolate the HDMECs according to previously published protocols21. Briefly, after separation of dermis from epidermis, incubate the dermis in trypsin for 40 min at 37 &#176;C and 5% CO2. After incubation time, transfer the dermis to a Petri dish containing culture medium, and scratch each piece with the use of a scalpel. Transfer the obtained cell suspension to a 50 mL plastic tube and centrifuge for 5 min at 300 x g and 22 &#176;C. Count the cell suspension using a Neubauer chamber and seed at a density of 1.2 x 104 cells/cm2. Change the medium completely every other day.</li>
	</ol>
	</li>
</ol>
2. Set-up and sterilization of spinner flasks, RCP microcarriers, and bioreactor
<ol>
	<li>Spinner flasks and RCP microcarriers
	<ol>
		<li>One day before the experiments, set-up the spinner flasks and prepare them for sterilization by autoclave. 
		NOTE: Leave the caps of the spinner flasks loose and, when possible, sterilize them in an upright position.
		<ol>
			<li>Wrap the spinner flasks with aluminum foil around the side caps separately in order to avoid contamination during unwrapping. Wrap the flasks with 1 - 2 layers of aluminum foil for autoclaving in order to minimize contamination risks after sterilization.</li>
		</ol>
		</li>
		<li>Weigh the required amount of RCP macroporous microcarriers in 20 mL of DPBS. 
		NOTE: 30 mg is required per spinner flask. Let them hydrate for at least 1 h before heat sterilization by autoclave (121 &#176;C for 15 min).</li>
	</ol>
	</li>
	<li>Perfusion bioreactor
	<ol>
		<li>Use the bioreactor system described to produce vascularized tissue equivalents20. This bioreactor consists of a vessel, in which the biomatrix is placed, a medium reservoir and a pressure bottle.
		<ol>
			<li>Connect the vessel with the medium reservoir and the pressure bottle through silicon tubes (see Figure 3C). Leave the caps loose and place it in an autoclave plastic bag, close well, and place this in a second autoclave plastic bag. Sterilize by autoclave.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
3. Culture of hBMSCs and HDMECs on RCP macroporous microcarriers
NOTE: The HDMECs used to seed the RCP microcarriers were labeled with RFP by lentiviral transduction. This protocol was modified after a previously published protocol5.
<ol>
	<li>Let the RCP macroporous microcarriers settle to the bottom and wash them with 10 mL of culture medium. Repeat 3 times.</li>
	<li>Wash spinner flasks with 10 mL of culture medium.</li>
	<li>Add 30 mg of RCP macroporous microcarriers per spinner flask in 8 mL of culture medium. Equilibrate the spinner flasks containing the RCP microcarriers at 37 &#176;C and 5% CO2 for 30 min, before seeding the cells. 
	NOTE: This is based on approximate dry weight of RCP microcarriers before sterilization.</li>
	<li>Seed 5 x 105 hBMSCs or HDMECs (passage 3) per spinner flask in 2 mL of culture medium.</li>
	<li>Place the spinner flasks on the stirrer plate and start stirring at 90 rpm for 5 min and rest for 55 min, at 37 &#176;C and 5% CO2, for a total of 1 h static/dynamic incubation. Repeat 4 times.</li>
	<li>Add 10 mL of cell culture medium per spinner flask and then start continuous stirring at 95 rpm. Incubate at 37 &#176;C and 5% CO2.</li>
	<li>With the use of a micropipette take samples at days 1, 4 and 7: 333 &#181;L for DNA content (~ 0.5 mg), 1000&#160;&#181;L for SEM analysis (~ 1.5 mg) and 333 &#181;L for live/dead staining (~ 0.5 mg). 
	NOTE: This is based on approximate dry weight of RCP microcarriers before sterilization.</li>
	<li>Change 10 mL of culture medium every other day. For this, wait until the RCP microcarriers settle to the bottom and very carefully aspirate 10 mL from the spinner flask, with the use of a 10 mL pipette, and then add 10 mL of fresh culture medium.</li>
	<li>Keep the cultures on the stirrer plate set at 95 rpm, at 37 &#176;C and 5% CO2 for 7 days.</li>
</ol>
4. DNA content, SEM analysis, live dead staining, and sprouting assay
<ol>
	<li>DNA content
	<ol>
		<li>With the use of a micropipette, collect approx. 0.5 mg of microcarriers from each spinner flask (contained in 333 &#181;L of suspension) and transfer to a 2 ml plastic tube. 
		NOTE: This is based on approximate dry weight of RCP microcarriers before sterilization.</li>
		<li>Wash once with 1 mL of DPBS, then aspirate supernatant.</li>
		<li>Remove remaining DPBS in a lyophilizer and weight the lyophilized microcarriers for later normalization.</li>
		<li>Freeze samples at -80 &#176;C for at least 6 h.</li>
		<li>Incubate samples at 60 &#176;C overnight with 300 &#181;L papain 5 U/mL.</li>
		<li>Establish the appropriate titration curves following the manufacturer&#180;s instructions of the quantitation assay (e.g., PicoGreen dsDNA assay) kit and perform measurement of the fluorescent signal at 520 nm with a luminescence detector.</li>
	</ol>
	</li>
	<li>Scanning electron microscopy (SEM) analysis
	<ol>
		<li>Collect ca. 1 mg of microcarriers from each spinner flask and transfer to a 2 mL plastic tube.</li>
		<li>Wash microcarriers once with 1 mL of DPBS.</li>
		<li>Remove DPBS and incubate in 1 mL of 70% ethanol for 1 h.</li>
		<li>Remove supernatant and incubate in 1 mL of 80% ethanol for 1 h.</li>
		<li>Remove supernatant and incubate in 1 mL of 90% ethanol for 1 h.</li>
		<li>Remove supernatant and incubate in 1 mL of 100% ethanol for 1 h.</li>
		<li>Remove supernatant and incubate in 1 mL of hexamethyldisilazane for 10 min.</li>
		<li>Open cap and let the samples dry overnight under a chemical hood.</li>
		<li>Fix samples on appropriate support and proceed to SEM analysis according to established protocols.</li>
	</ol>
	</li>
	<li>DAPI and Live/Dead staining
	<ol>
		<li>At day 2, 4, and 7 after cell seeding on the microcarriers collect 0.5 mg of microcarriers from each spinner flask and transfer to a 2 mL plastic tube. Wash once with 1 mL of DPBS.</li>
		<li>DAPI
		<ol>
			<li>Fix samples for 10 min in enough 4% paraformaldehyde to cover them.</li>
			<li>Wash once with 1 mL of DPBS.</li>
			<li>Incubate at room temperature with the staining solution for 45 min on a rocking shaker.</li>
			<li>Detect DAPI fluorescent signal on a fluorescence microscope with a 420 nm emission filter and acquire images.</li>
		</ol>
		</li>
		<li>Live/Dead Staining
		<ol>
			<li>Incubate samples in 500 &#181;L of Live/Dead dye solution. Keep the samples at 37 &#176;C for 20 min, protected from light.</li>
			<li>Wash once with 1 mL of DPBS.</li>
			<li>Detect fluorescence signal of living cells with a 490 nm emission filter and of dead cells with a 545 nm emission filter on a fluorescence microscope. Acquire images.</li>
		</ol>
		</li>
		<li>Sprouting assay for HDMECs
		<ol>
			<li>Mix 330 &#181;L of collagen R solution 0.4%, 170 &#181;L of 0.1% acetic acid, and 50 &#181;L of medium M199 in a 15 mL plastic tube. Keep on ice.</li>
			<li>Take ~0.375 mg of microcarriers (250 &#181;L) with the use of a micropipette. Transfer to a 1.5 mL plastic tube, wait for the RCP microcarriers to settle to the bottom and remove the medium completely. 
			NOTE: This is based on approximate dry weight of RCP microcarriers before sterilization.</li>
			<li>Add the volume of NaOH 0.2 N required to neutralize the collagen mixture and immediately mix it with the RCP microcarriers. 
			NOTE: Determine the required amount of NaOH 0.2 N in advance by adding it to the mixture until change in pH (turn from yellow color to pink), and then placing the mixture in the incubator for 1 min, after which a gel must be formed.</li>
			<li>Plate the collagen gel-RCP microcarriers mixture in a well of a 24-well plate. Incubate at 37 &#176;C and 5% CO2 for 30 min.</li>
			<li>Add 200 &#181;L of the vascular endothelial growth factor (e.g., VascuLife VEGF-Mv culture medium). Incubate at 37 &#176;C and 5% CO2 for 24 h.</li>
			<li>Observe under a inversed phase contrast microscope, using a 10X magnification object, and acquire images.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
5. Biomatrix seeding and start of bioreactor system for HDMECs
<ol>
	<li>One day before starting the bioreactor cultures, cut open BioVaSc (the biomatrix) longitudinally on the antimesenteric side and fix it in a previously disinfected polycarbonate frame (see Figure 3A, B). 
	NOTE: The BioVaSc is a biomatrix obtained from a decellularized porcine jejunal segment, prepared as previously published21. 
	NOTE: As the polycarbonate frame cannot be sterilized by heat, disinfect it by 20 min incubation in sonic bath (40 kHz) followed by 3 times incubation in ethanol 70% for 25 min. Then, wash the frame 3 times with DPBS sterile.
	<ol>
		<li>Place the biomatrix-frame system in a 100 mm Petri dish and fill it with the vascular endothelial growth factor + 1% Penicillin Streptomycin (Pen/Strep).</li>
		<li>Equilibrate the biomatrix by incubating overnight at 37 &#176;C and 5% CO2.</li>
	</ol>
	</li>
	<li>After detaching and counting the cells, inject 10 x 107 HDMECs in 1000 &#181;L of culture medium through the preserved vasculature of the biomatrix, with the use of a 2 mL syringe. 
	NOTE: Inject ~700 &#181;L through the arterial inlet and ~300 &#181;L through the vein outlet.</li>
	<li>Incubate at 37 &#176;C and 5% CO2 for 3 h, to allow cell attachment.</li>
	<li>Fill the bioreactor system with 350 mL of the vascular endothelial growth factor + 1% Pen/Strep and incubate at 37 &#176;C and 5% CO2.</li>
	<li>Place the frame inside the vessel of the bioreactor. Connect the arterial and venous pedicles to the vessel.</li>
	<li>Start perfusion by connecting the bioreactor system to a Peristaltic pump. Set the pressure at 10 mmHg and amplitude &#177;1.</li>
	<li>Increase the pressure stepwise every hour until reaching 100 mmHg of pressure, amplitude &#177;20.</li>
	<li>Keep the bioreactor system for 7 days under these conditions at 37 &#176;C and 5% CO2.</li>
</ol>
6. Addition of RFP-HDMECs to the biomatrix
<ol>
	<li>Mix 330 &#181;L of 0.4% collagen R solution, 170 &#181;L of 0.1% acetic acid, and 50 &#181;L of medium M199 in a 15 mL Falcon tube. Keep on ice.</li>
	<li>Take the RCP microcarriers from the spinner flasks. Remove the culture medium completely.</li>
	<li>Add the volume of NaOH 0.2 N required to neutralize the collagen mixture and immediately mix it with the RCP microcarriers.</li>
	<li>Add the collagen gel-RCP microcarriers mixture on the lumen of the biomatrix. Allow gelation for 30 min.</li>
	<li>In parallel, change the medium of the bioreactor by removing the medium completely and then add 350 mL of fresh, pre warmed, vascular endothelial growth factor + 1% Pen/Strep.</li>
	<li>Connect the bioreactor to the Peristaltic pump and set up the pressure at 100 mmHg and amplitude &#177;20.</li>
	<li>Change the medium every 7 days.</li>
	<li>Keep the bioreactor in culture for 21 days.</li>
</ol>
7. Analysis and read-outs of bioreactor cultures
<ol>
	<li>Preparation of the samples
	<ol>
		<li>Disconnect the biomatrix from the bioreactor and remove it from the polycarbonate frame.</li>
		<li>Cut the obtained piece into 6 sections: 2 for MTT, 2 for paraffin embedding/staining and 2 for SEM analysis.</li>
	</ol>
	</li>
	<li>Paraffin embedding
	<ol>
		<li>Place the sections in a Petri dish and wash with enough DPBS to cover them. Discard the DPBS.</li>
		<li>Fix the sections in 4% paraformaldehyde overnight.</li>
		<li>Prepare the cassettes for paraffin embedding and perform it according to standard protocols.</li>
		<li>Prepare the sections in paraffin blocks and cut samples of 3 &#181;m for H&#38;E staining, with the use of a microtome.</li>
	</ol>
	</li>
	<li>H&#38;E staining
	<ol>
		<li>Stain rehydrated sections according to standard protocols32.</li>
	</ol>
	</li>
	<li>Scanning electron microscopy (SEM) analysis
	<ol>
		<li>Place the sections in a Petri dish and wash with enough DPBS to cover them. Discard the DPBS.</li>
		<li>Fix the sections with 4.75% glutaraldehyde&#160;overnight.</li>
		<li>Perform dehydration chain with increasing concentrations of ethanol 50% - 100%.</li>
		<li>Cover the sections with hexamethyldisilazane (HMDS) for 10 min. Discard the used HMDS and add fresh HMDS.</li>
		<li>Allow the sections to dry overnight under fume hood, and then prepare the samples for imaging.</li>
	</ol>
	</li>
	<li>3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT)
	<ol>
		<li>Mix the MTT reagent with the vascular endothelial growth factor, in a concentration of 1 mg/mL.</li>
		<li>Incubate the sections in the MTT solution for 90 min at 37 &#176;C and 5% CO2.</li>
		<li>Observe the vascular structures and cell-seeded microcarriers macroscopically or under a bright-field microscope, to detect metabolically active cells. Acquire images.</li>
	</ol>
	</li>
</ol>

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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+medium+perfusion+on+cell-seeded+three-dimensional+bone+constructs+in+vitro.">Effects of medium perfusion on cell-seeded three-dimensional bone constructs in vitro.</a> Tissue Engineering. 9 (6), 1197-1203 (2004).</li><li>Schanz, J., Pusch, J., Hansmann, J., Walles, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vascularised+human+tissue+models:+a+new+approach+for+the+refinement+of+biomedical+research.">Vascularised human tissue models: a new approach for the refinement of biomedical research.</a> Journal of biotechnology. 148 (1), 56-63 (2010).</li><li>Steinke, M., Gross, R., Walles, H., Sch&#252;tze, K., Walles, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+engineered+3D+human+airway+mucosa+model+based+on+a+SIS+scaffold.">An engineered 3D human airway mucosa model based on a SIS scaffold.</a> Biomaterials. 35 (26), 7355-7362 (2014).</li><li>Moll, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+Engineering+of+a+Human+3D+in+vitro+Tumor+Test+System.">Tissue Engineering of a Human 3D in vitro Tumor Test System.</a> J. Vis. Exp. (78), e50460 (2013).</li><li>Groeber, F., Kahlig, A., Loff, S., Walles, H., Hansmann, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+bioreactor+system+for+interfacial+culture+and+physiological+perfusion+of+vascularized+tissue+equivalents.">A bioreactor system for interfacial culture and physiological perfusion of vascularized tissue equivalents.</a> Biotechnology Journal. 8, 308-316 (2013).</li><li>Alberts, B., Johnson, A., Lewis, J., Raff, M., Roberts, K., Walter, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Blood+vessels+and+endothelial+cells.">Blood vessels and endothelial cells.</a> Molecular Biology of the Cells. , (2002).</li><li>Logsdon, E. A., Finley, S. D., Popel, A. S., Gabhann, F. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+systems+biology+view+of+blood+vessel+growth+and+remodeling.">A systems biology view of blood vessel growth and remodeling.</a> Journal of Cellular and Molecular Medicine. 18 (8), 1491-1508 (2014).</li><li>Scheller, K., Dally, I., Hartmann, N., M&#252;nst, B., Braspenning, J., Walles, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Upcyte&#174;+Microvascular+endothelial+cells+repopulate+decellularized+scaffold.">Upcyte&#174; Microvascular endothelial cells repopulate decellularized scaffold.</a> Tissue Engineering Part C: Methods. 19 (1), 57-67 (2012).</li><li>Nietzer, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mimicking+Metastases+Including+Tumor+Stroma:+A+New+Technique+to+Generate+a+Three-Dimensional+Colorectal+Cancer+Model+Based+on+a+Biological+Decellularized+Intestinal+Scaffold.">Mimicking Metastases Including Tumor Stroma: A New Technique to Generate a Three-Dimensional Colorectal Cancer Model Based on a Biological Decellularized Intestinal Scaffold.</a> Tissue Engineering Part C: Methods. 22 (7), 621-635 (2016).</li><li>G&#246;ttlich, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Combined+3D+Tissue+Engineered+In+Vitro/In+Silico+Lung+Tumor+Model+for+Predicting+Drug+Effectiveness+in+Specific+Mutational+Backgrounds.">A Combined 3D Tissue Engineered In Vitro/In Silico Lung Tumor Model for Predicting Drug Effectiveness in Specific Mutational Backgrounds.</a> J. Vis. Exp. (110), e53885 (2016).</li><li>Stratmann, A. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment+of+a+human+3D+lung+cancer+model+based+on+a+biological+tissue+matrix+combined+with+a+Boolean+in+silico+model.">Establishment of a human 3D lung cancer model based on a biological tissue matrix combined with a Boolean in silico model.</a> Molecular oncology. 8 (2), 351-365 (2014).</li><li>Groeber, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+first+vascularized+skin+equivalent+for+as+an+alternative+to+animal+experimentation.">A first vascularized skin equivalent for as an alternative to animal experimentation.</a> Altex. 33 (4), 415-422 (2016).</li><li>Plunkett, N., O'Brien, F. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bioreactors+in+tissue+engineering.">Bioreactors in tissue engineering.</a> Technology and Health Care. 19 (1), 55-69 (2011).</li><li>Nienow, A. W., Rafiq, Q. A., Coopman, K., Hewitt, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+potentially+scalable+method+for+the+harvesting+of+hMSCs+from+microcarriers.">A potentially scalable method for the harvesting of hMSCs from microcarriers.</a> Biochemical Engineering Journal. 85, 79-88 (2014).</li><li>Fischer, A. H., Jacobson, K. A., Rose, J., Zeller, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hematoxylin+and+eosin+staining+of+tissue+and+cell+sections.">Hematoxylin and eosin staining of tissue and cell sections.</a> Cold Spring Harbor Protocols. 2008 (5), pdb-prot4986 (2008).</li></ol>

The authors have nothing to disclose.

One main goal of microcarrier is the expansion of cells while maintaining their differentiation in order to deliver cells to the place of need. The represented method introduce RCP microcarriers where cells were able to attach, proliferate, and colonize the microcarriers with high cell density. This was observed by live/dead staining, in which more than 90% of viable cells were detected while only few dead cells were obtained after 7 days of dynamic cultures. Likewise, the SEM images confirmed that the cells covered the entire surface of the microcarriers after 7 days of cultures.
To ensure cell survival in 3D models, it is important to maintain the supply of oxygen and nutrients to the cells and allow removal of waste substances. Blood vessels are the responsible structures of this exchange process, in which endothelial cells play a key role since they are the cells lining the inner part of blood vessels23. They have the capability to proliferate, migrate, adhere, sprout, and form vessel-like structures24. These properties were maintained after culture of RFP-HDMECs on the RCP microcarriers, since it was observed that the cells were able to adopt the sprouting phenotype (see Figure 2). We could prove here that these colonized RCP microcarriers were effectively delivering primary endothelial cells to re-seed the lumen of former vessels of the BioVaSc biomatrix. This suggests our system to be suitable to improve the vascularization of tissue engineered implants for clinical applications. Derivatives of this matrix have been successfully used in vascularization approaches25, as well as in in vitro tumor test sytems21,26,27,28 and for the production of skin equivalents as alternatives to animal experimentation29. Here it is used in an open, flattened set-up and placed in a perfusion bioreactor as applied for the generation of tissue equivalents22.
Bioreactors have been used in tissue engineering to produce tissue equivalents that could help ease the demand of organs while reducing the associated risks like rejection and morbidity30. However, producing tissue-like constructs is a very complex process that involves the use of tissue-specific cell types, a suitable scaffold and appropriate growth factors that allow the proper differentiation and assembling into 3D tissues. One major drawback of the engineered constructs is the lack of a proper vascular network that supports cells survival both in vitro and in vivo17. Here we combine the RCP-colonized microcarriers with a decellularized matrix in a bioreactor model, providing the cell type required, a scaffold that allows cell adhesion and vessel formation and a specific culture medium that provides the essential factors for the cells to proliferate and maintain their functional properties. A significant result of this model is the ability of the HDMECs to migrate from the RCP microcarriers to the scaffold without any additional detachment or digestion as necessary in other approaches31. Afterwards, they colonized specifically the vascular structures of the matrix, proving the concept that macroporous microcarriers can be used as cell delivery system for Regenerative Medicine purposes.
Altogether, this protocol represents a promising strategy in regenerative medicine for obtaining a maximized cell expansion and it could serve as cell delivery to implantation sites, where it could improve vascularization support for tissue engineered grafts.

One general problem in tissue engineering applications is to yield a high cell mass with the correct differentiation phenotype at the location of need. The application of microcarriers to address this issue started in 1967 with increasing significance to date in fields such as orthopaedic tissue engineering for large-scale generation of skin, bone, cartilage, and tendons1. They allow the handling of adherent cultures in ways similar to that of suspension cultures2 by expanding cells on microscale three-dimensional (3D) substrates. Thereby cells experience a homogeneous nutrient supply and cell-matrix interactions that lead to better maintenance of in vivo3,4 differentiation which is often lost over time in 2D approaches5. A higher surface-to-volume ratio - eventually leading to higher cell yields6,7, higher gas and nutrient exchange rates comparing to static systems8, the possibility to regulate and subject the culture to physical stimuli9, and the potential for scaling up of the expansion process7 are further advantages. Several features such as diameter, density, porosity, surface charge, and adhesion properties10,11 distinguish the different commercially available micro- and macro-carriers. However, one of the main advantage is their delivery potential as microtissues to site defect or demand.
For applications of the microcarrier technology in bone tissue engineering, we illustrated in a previous report12 the production of a new microcarrier type constituted of a recombinant collagen I peptide (RCP, commercially available as Cellnest). This new microcarrier allows the GMP-compliant up scaling of scaffold and cell production, as needed for cell delivery in a clinical scenario. In this context, tuning of scaffold stability, degradation rate, and surface properties through proper choice of a suited crosslinking strategy allows to adapt the technique to the selected application, cell type of interest or target tissue13. In particular, the potential employment of this microcarrier as an injectable cell delivery system for therapeutic application14 makes them particularly interesting in a clinical setting.
In this paper, we therefore illustrate the culturing procedure for the isolation and expansion of human bone marrow-derived mesenchymal stromal cells (hBMSCs) and human dermal microvascular endothelial cells (HDMECs) on collagen-I-based recombinant peptide-based microcarriers, and their preparation for delivery in a clinical setting. Furthermore, we describe additional protocols useful for the maintenance of cell viability upon implantation.
Cell viability after implantation is in fact strongly dependent on vascularization15,16,17, which ensures exchange of oxygen and nutrients and facilitates waste removal. Bioreactors constitute one approach to overcome vascularization challenges in tissue engineering and maintain cell viability, through perfusion of culture medium providing thereby oxygen and nutrients18. Here, we illustrate an in vitro method to evaluate the migration capability of microvascular endothelial cells from the RCP microcarriers to a biomatrix and their ability to contribute to the de novo vascularization and angiogenesis. This biomatrix is a decellularized segment of porcine jejunum termed BioVaSc (Biological Vascularized Scaffold), rich in collagen and elastin and with preserved vascular structures, which includes a feeding artery and a draining vein19 that has been applied for implantation issues20.

<table><tbody><tr><td>3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide (MTT)</td><td>Serva Electrophoresis GmbH</td><td>20395.01</td><td></td></tr><tr><td>4&amp;rsquo;,6-Diamidino-2-phenylindoldihydrochloride (DAPI)</td><td>Sigma-Aldrich</td><td>D9542</td><td></td></tr><tr><td>Acetic acid 100%</td><td>Sigma-Aldrich</td><td>533,001</td><td></td></tr><tr><td>Analytical balance Kern EG 2200-2NM</td><td>Kern &amp;amp; Sohn GmbH</td><td></td><td></td></tr><tr><td>Ascorbate-2-phosphate</td><td>Sigma-Aldrich</td><td>A8960</td><td></td></tr><tr><td>Bioreactor</td><td>Chair of Tissue Engineering and Regenerative Medicine, Wuerzburg, Germany</td><td></td><td></td></tr><tr><td>Bright field microscope Axiovert 40C</td><td>Carl Zeiss AG</td><td></td><td></td></tr><tr><td>Cellnest</td><td>Fujifilm</td><td></td><td></td></tr><tr><td>Centrifuge tubes (15 mL, 50 mL)</td><td>Greiner Bio-One</td><td></td><td></td></tr><tr><td>Collagen R solution 0,4%</td><td>Serva Electrophoresis GmbH</td><td>47254.01</td><td></td></tr><tr><td>DMEM-F12</td><td>Gibco</td><td>11320-033</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Saline</td><td>Sigma-Aldrich</td><td>D8537</td><td>Modified, without calcium chloride and magnesium chloride</td></tr><tr><td>Eosin 1%</td><td>Morphisto</td><td>10177.01000</td><td></td></tr><tr><td>Ethanol 96%</td><td>Carl Roth GmbH</td><td>T171.4</td><td>Denatured</td></tr><tr><td>Fetal calf serum (FCS)</td><td>Bio&amp;amp;SELL</td><td>FCS.ADD.0500</td><td>not heat-inactivated</td></tr><tr><td>Fluorescence microscope BZ-9000</td><td>Keyence</td><td></td><td></td></tr><tr><td>Haematoxylin</td><td>Morphisto</td><td>10231.01000</td><td></td></tr><tr><td>Hexamethyldisilazane</td><td>Sigma-Aldrich</td><td>440191</td><td>Reagent grade, &amp;ge;99%</td></tr><tr><td>Incubator for bioreactor</td><td>Chair of Tissue Engineering and Regenerative Medicine, Wuerzburg, Germany</td><td></td><td></td></tr><tr><td>Live/Dead Cell Double Staining Kit</td><td>Fluka</td><td>04511KT-F</td><td></td></tr><tr><td>Magnetic stirrer plate</td><td>2Mag</td><td>80002</td><td></td></tr><tr><td>Medium 199</td><td>Sigma-Aldrich</td><td>M0650</td><td>10X</td></tr><tr><td>Microplate reader&lt;br/&gt; Tecan Infinite M200</td><td>Tecan</td><td></td><td></td></tr><tr><td>Needle 21G 16mm</td><td>VWR</td><td>613-5389</td><td></td></tr><tr><td>Papain from papaya latex</td><td>Sigma-Aldrich</td><td>P4762</td><td>lyophilized powder, &amp;ge; 10 units/mg protein</td></tr><tr><td>Paraffin</td><td>Carl Roth GmbH</td><td>6642.6</td><td></td></tr><tr><td>Penicillin/Streptomycin</td><td>Sigma-Aldrich</td><td>P4333</td><td></td></tr><tr><td>Peristaltic pump</td><td>Ismatec</td><td></td><td></td></tr><tr><td>Quanti-iT PicoGreen dsDNA assay kit</td><td>Thermo Fischer Scientific</td><td>P7589</td><td></td></tr><tr><td>Histofix 4%</td><td>Carl Roth GmbH</td><td>P087</td><td></td></tr><tr><td>Scanning Electron Microscope Supra 25</td><td>Carl Zeiss AG</td><td></td><td></td></tr><tr><td>Sodium hydroxide solution 1.0 N</td><td>Sigma-Aldrich</td><td>S2770</td><td></td></tr><tr><td>Spinner flasks (25 mL)</td><td>Wheaton</td><td>356879</td><td></td></tr><tr><td>Syringe 1 mL</td><td>VWR</td><td>720-2561</td><td></td></tr><tr><td>Tissue culture flasks (25 cm&lt;sup&gt;2&lt;/sup&gt;, 75 cm&lt;sup&gt;2&lt;/sup&gt;, 150 cm&lt;sup&gt;2&lt;/sup&gt;)</td><td>TPP Techno Plastik Products AG</td><td></td><td></td></tr><tr><td>Trypan blue 0.4%</td><td>Sigma-Aldrich</td><td>T8154</td><td></td></tr><tr><td>VascuLife VEGF-Mv</td><td>Lifeline cell technology</td><td>LL-0005</td><td></td></tr></tbody></table>

recombinant collagen i peptide microcarriers for cell expansion and their potential use as cell delivery system in a bioreactor model

Tissue engineering is a promising field, focused on developing solutions for the increasing demand on tissues and organs regarding transplantation purposes. The process to generate such tissues is complex, and includes an appropriate combination of specific cell types, scaffolds, and physical or biochemical stimuli to guide cell growth and differentiation. Microcarriers represent an appealing tool to expand cells in a three-dimensional (3D) microenvironment, since they provide higher surface-to volume ratios and mimic more closely the in vivo situation compared to traditional two-dimensional methods. The vascular system, supplying oxygen and nutrients to the cells and ensuring waste removal, constitutes an important building block when generating engineered tissues. In fact, most constructs fail after being implanted due to lacking vascular support. In this study, we present a protocol for endothelial cell expansion on recombinant collagen-based microcarriers under dynamic conditions in spinner flask and bioreactors, and we explain how to determine in this setting cell viability and functionality. In addition, we propose a method for cell delivery for vascularization purposes without additional detachment steps necessary. Furthermore, we provide a strategy to evaluate the cell vascularization potential in a perfusion bioreactor on a decellularized&#160;biological matrix. We believe that the use of the presented methods could lead to the development of new cell-based therapies for a large range of tissue engineering applications in the clinical practice.

As shown in Figure 1A, we obtained high number of viable cells on the RCP microcarriers after 7 days of culture, determined by live/dead staining. Those results were confirmed by SEM analysis, in which completely colonized microcarriers were observed around the pores, partly overgrowing them (Figure 1B). On the other hand, experiments in which cells were not evenly seeded resulted in several empty microcarriers. Failed experiments are characterized by an abnormally high number of dead cells that should not be over 10% of the total cell amount (Figure 1C). Moreover, DNA content quantification of HDMECs showed an increase of dsDNA over time (Figure 1D).
Additionally, RFP-HDMECs were able to maintain their functionality after culture on RCP microcarriers, as shown in Figure 2, where the cells adopted a sprouting phenotype when cultured in a collagen gel.
Furthermore, we used RFP-HDMECs-colonized RCP microcarriers to re-seed the lumen of the biomatrix BioVaSc. For this, we employed a bioreactor system for physiological perfusion of vascularized tissue equivalents22 (Figure 3C) in which we cut-open the biomatrix and placed it in a polycarbonate frame, so that the lumen was exposed in an even set-up (Figure 3B).
After 21 days of bioreactor cultures, metabolically active cells were present both on the preserved vasculature of the biomatrix as well as on the RCP microcarriers (Figure 4A and Figure 3B). Failed experiments or a wrong choice of slices during sectioning of the histological samples can lead to absence of colonizing endothelial cells in the lumen of the vessels of the biomatrix or on the microcarriers. These results could be confirmed by SEM analysis of the biomatrix sections, where it could be observed that some areas of the RCP microcarriers were still covered by cells and that the RCP microcarriers were in close proximity to the biomatrix (Figure 4C and 4D).
Finally, H&#38;E staining confirmed the presence of HDMECs on the biomatrix, specifically in the vascular structures (Figure 5A). When observed under fluorescence microscope, the cells colonizing the vascular structures of the biomatrix resulted to be RFP-HDMECs (Figure 5B), indicating the migration of the cells from the RCP microcarriers to the biomatrix. Some RFP-HDMECs were also observed on the RCP microcarriers (Figure 5C and 5D). In experiments in which cells did not survived due to altered culturing conditions (e.g. shorter culture times), they could not be detected inside the vascular structure with none of the previously reported techniques (Figure 5E and 5F).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57363/57363fig1.jpg" /> 
Figure 1: Culture of HDMECs and hBMSCs on RCP microcarriers. (A, C) live/dead staining after 7 days of dynamic cultures. (B) SEM analysis of HDMECs cultured on RCP microcarriers after 7 days of dynamic cultures. (D) DNA content determined by PicoGreen assay kit. Data expressed as mean &#177; standard deviation (n=3). Scale bars: 200 &#181;m for A, 88 &#181;m for B, and 100 &#181;m for C. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57363/57363fig1large.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/57363/57363fig2.jpg" /> 
Figure 2: Sprouting assay. Sprouts of RFP-HDMECs can be observed under bright-field (A) and fluorescence microscopy (B), emerging from the colonized RCP microcarrier after 24 h of culture in a collagen gel. (C) Overlay image. Scale bars: 100 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57363/57363fig2large.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/57363/57363fig3.jpg" /> 
Figure 3: Biomatrix and bioreactor set-up. (A) Preparation of the biomatrix and mounting on the polycarbonate frame (B). (C) Bioreactor set-up. The vessel is connected to the medium reservoir and the pressure bottle through silicon tubes, and the whole system is connected to a peristaltic pump. The pressure is measured through a pressure sensor. AI: arterial inlet; VO: venous outlet. This figure has been modified from Groeber et al.22 <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57363/57363fig3large.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/57363/57363fig4.jpg" /> 
Figure 4: Bioreactor read-out. MTT assay performed on a biomatrix section after 21 days of culture, in which metabolically active cells were observed in the preserved vasculature of the biomatrix (A) as well as on the RCP microcarriers (B). (C, D) SEM images of RCP microcarriers on a biomatrix section. Scale bars: 0.28 cm for A, 0.45 cm for B, 47 &#181;m for C, and 225 &#181;m for D. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57363/57363fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57363/57363fig5.jpg" /> 
Figure 5: H&#38;E staining &#38; fluorescence images. (A, C, E) H&#38;E staining. (B, D, F)&#160;After 21 days of culture,&#160;RFP-HDMECs were observed inside of the vasculature of the biomatrix (arrows) as well as on the RCP microcarriers (arrow heads). Scale bars: 50 &#181;m for A-D and 30 &#181;m for E and F. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57363/57363fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Recombinant Collagen I Peptide Microcarriers for Cell Expansion and Their Potential Use As Cell Delivery System in a Bioreactor Model at JoVE.com

Recombinant Collagen I Peptide Microcarriers for Cell Expansion and Their Potential Use As Cell Delivery System in a Bioreactor Model

We propose a cell expansion protocol on macroporous microcarriers and their use as delivery system in a perfusion bioreactor to seed a decellularized tissue matrix. We also include different techniques to determine cell proliferation and viability of cells cultured on microcarriers. Furthermore, we demonstrate functionality of cells after bioreactor cultures.

Tissue engineering is a promising field, focused on developing solutions for the increasing demand on tissues and organs regarding transplantation ...

recombinant-collagen-i-peptide-microcarriers-for-cell-expansion-their

Nanyang Technological University

Research

JoVE Journal

Bioengineering

7.5K Views.  University Hospital Wuerzburg. Wij stellen voor een cel expansie protocol betreffende macroporeuze microcarriers en hun gebruik als levering systeem in een bioreactor perfusie om het zonebeheer van een matrix decellularized weefsel. Wij omvatten ook verschillende technieken om te bepalen van celproliferatie en levensvatbaarheid van cellen gekweekt op microcarriers. Bovendien tonen we functionaliteit van cellen na bioreactor culturen.

Video: Recombinant Collagen I Peptide Microcarriers for Cell Expansion and Their Potential Use As Cell Delivery System in a Bioreactor Model

Constructing a Collagen Hydrogel for the Delivery of Stem Cell-loaded Chitosan Microspheres

Multipotent stem cells have been shown to be extremely useful in the field of regenerative medicine1-3. However, in order to use these cells effectively for tissue regeneration, a number of variables must be taken into account. These variables include: the total volume and surface area of the implantation site, the mechanical properties of the tissue and the tissue microenvironment, which includes the amount of vascularization and the components of the extracellular matrix. Therefore, the materials being used to deliver these cells must be biocompatible with a defined chemical composition while maintaining a mechanical strength that mimics the host tissue. These materials must also be permeable to oxygen and nutrients to provide a favorable microenvironment for cells to attach and proliferate. Chitosan, a cationic polysaccharide with excellent biocompatibility, can be easily chemically modified and has a high affinity to bind with in vivo macromolecules4-5. Chitosan mimics the glycosaminoglycan portion of the extracellular matrix, enabling it to function as a substrate for cell adhesion, migration and proliferation. In this study we utilize chitosan in the form of microspheres to deliver adipose-derived stem cells (ASC) into a collagen based three-dimensional scaffold6. An ideal cell-to-microsphere ratio was determined with respect to incubation time and cell density to achieve maximum number of cells that could be loaded. Once ASC are seeded onto the chitosan microspheres (CSM), they are embedded in a collagen scaffold and can be maintained in culture for extended periods. In summary, this study provides a method to precisely deliver stem cells within a three dimensional biomaterial scaffold.

A major hurdle in current stem cell therapies is determining the most effective method to deliver these cells to host tissues. Here, we describe a chitosan-based delivery method that is efficient and simple in approach, while allowing adipose-derived stem cells to maintain their multipotency.

Multipotent stem cells have been shown to be extremely useful in the field of regenerative medicine1-3. However, in order to use these cells effectively ...

Engineering 3D Cellularized Collagen Gels for Vascular Tissue Regeneration

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. Collagen has been extensively used for a wide range of biomedical applications and is considered a valid alternative to synthetic materials due to its inherent biocompatibility (i.e., low antigenicity, inflammation, and cytotoxic responses). However, the limited mechanical properties and the related low hand-ability of collagen gels have hampered their use as scaffold materials for vascular tissue engineering. Therefore, the rationale behind this work was first to engineer cellularized collagen gels into a tubular-shaped geometry and second to enhance smooth muscle cells driven reorganization of collagen matrix to obtain tissues stiff enough to be handled.
The strategy described here is based on the direct assembling of collagen and smooth muscle cells (construct) in a 3D cylindrical geometry with the use of a molding technique. This process requires a maturation period, during which the constructs are cultured in a bioreactor under static conditions (without applied external dynamic mechanical constraints) for 1 or 2 weeks. The &#8220;static bioreactor&#8221; provides a monitored and controlled sterile environment (pH, temperature, gas exchange, nutrient supply and waste removal) to the constructs. During culture period, thickness measurements were performed to evaluate the cells-driven remodeling of the collagen matrix, and glucose consumption and lactate production rates were measured to monitor the cells metabolic activity. Finally, mechanical and viscoelastic properties were assessed for the resulting tubular constructs. To this end, specific protocols and a focused know-how (manipulation, gripping, working in hydrated environment, and so on) were developed to characterize the engineered tissues.

In this work, we present a technique for the rapid fabrication of living vascular tissues by direct culturing of collagen, smooth muscle cells and endothelial cells. In addition, a new protocol for the mechanical characterization of engineered vascular tissues is described.

Synthetic materials are known to initiate clinical complications such as inflammation, stenosis, and infections when implanted as vascular substitutes. ...

Minced Tissue in Compressed Collagen: A Cell-containing Biotransplant for Single-staged Reconstructive Repair

Conventional techniques for cell expansion and transplantation of autologous cells for tissue engineering purposes can take place in specially equipped human cell culture facilities. These methods include isolation of cells in single cell suspension and several laborious and time-consuming events before transplantation back to the patient. Previous studies suggest that the body itself could be used as a bioreactor for cell expansion and regeneration of tissue in order to minimize ex vivo manipulations of tissues and cells before transplanting to the patient. The aim of this study was to demonstrate a method for tissue harvesting, isolation of continuous epithelium, mincing of the epithelium into small pieces and incorporating them into a three-layered biomaterial. The three-layered biomaterial then served as a delivery vehicle, to allow surgical handling, exchange of nutrition across the transplant, and a controlled degradation. The biomaterial consisted of two outer layers of collagen and a core of a mechanically stable and slowly degradable polymer. The minced epithelium was incorporated into one of the collagen layers before transplantation. By mincing the epithelial tissue into small pieces, the pieces could be spread and thereby the propagation of cells was stimulated. After the initial take of the transplants, cell expansion and reorganization would take place and extracellular matrix mature to allow ingrowth of capillaries and nerves and further maturation of the extracellular matrix. The technique minimizes ex vivo manipulations and allow cell harvesting, preparation of autograft, and transplantation to the patient as a simple one-stage intervention. In the future, tissue expansion could be initiated around a 3D mold inside the body itself, according to the specific needs of the patient. Additionally, the technique could be performed in an ordinary surgical setting without the need for sophisticated cell culturing facilities.

Tissue engineering often includes in vitro expansion in order to create autografts for tissue regeneration. In this study a method for tissue expansion, regeneration, and reconstruction in vivo was developed in order to minimize the processing of cells and biological materials outside the body.

Conventional techniques for cell expansion and transplantation of autologous cells for tissue engineering purposes can take place in specially equipped ...

Improving 2D and 3D Skin In Vitro Models Using Macromolecular Crowding

The glycoprotein family of collagens represents the main structural proteins in the human body, and are key components of biomaterials used in modern tissue engineering. A technical bottleneck is the deposition of collagen in vitro, as it is notoriously slow, resulting in sub-optimal formation of connective tissue and subsequent tissue cohesion, particularly in skin models. Here, we describe a method which involves the addition of differentially-sized sucrose co-polymers to skin cultures to generate macromolecular crowding (MMC), which results in a dramatic enhancement of collagen deposition. Particularly, dermal fibroblasts deposited a significant amount of collagen I/IV/VII and fibronectin under MMC in comparison to controls.
The protocol also describes a method to decellularize crowded cell layers, exposing significant amounts of extracellular matrix (ECM) which were retained on the culture surface as evidenced by immunocytochemistry. Total matrix mass and distribution pattern was studied using interference reflection microscopy. Interestingly, fibroblasts, keratinocytes and co-cultures produced cell-derived matrices (CDM) of varying composition and morphology. CDM could be used as &#34;bio-scaffolds&#34; for secondary cell seeding, where the current use of coatings or scaffolds, typically from xenogenic animal sources, can be avoided, thus moving towards more clinically relevant applications.
In addition, this protocol describes the application of MMC during the submerged phase of a 3D-organotypic skin co-culture model which was sufficient to enhance ECM deposition in the dermo-epidermal junction (DEJ), in particular, collagen VII, the major component of anchoring fibrils. Electron microscopy confirmed the presence of anchoring fibrils in cultures developed with MMC, as compared to controls. This is significant as anchoring fibrils tether the dermis to the epidermis, hence, having a pre-formed mature DEJ may benefit skin graft recipients in terms of graft stability and overall wound healing. Furthermore, culture time was condensed from 5 weeks to 3 weeks to obtain a mature construct, when using MMC, reducing costs.

Improving 2D and 3D Skin In Vitro Models Using Macromolecular Crowding

We present a protocol to obtain cell-derived matrices rich in extracellular matrix proteins, using macromolecular crowders (MMC). In addition, we present a protocol which incorporates MMC in 3D organotypic skin co-culture generation, which reduces culture time while maintaining maturity of construct.

The glycoprotein family of collagens represents the main structural proteins in the human body, and are key components of biomaterials used in modern ...

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

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

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

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

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

Fabrication of Extracellular Matrix-derived Foams and Microcarriers as Tissue-specific Cell Culture and Delivery Platforms

Cell function is mediated by interactions with the extracellular matrix (ECM), which has complex tissue-specific composition and architecture. The focus of this article is on the methods for fabricating ECM-derived porous foams and microcarriers for use as biologically-relevant substrates in advanced 3D in vitro cell culture models or as pro-regenerative scaffolds and cell delivery systems for tissue engineering and regenerative medicine. Using decellularized tissues or purified insoluble collagen as a starting material, the techniques can be applied to synthesize a broad array of tissue-specific bioscaffolds with customizable geometries. The approach involves mechanical processing and mild enzymatic digestion to yield an ECM suspension that is used to fabricate the three-dimensional foams or microcarriers through controlled freezing and lyophilization procedures. These pure ECM-derived scaffolds are highly porous, yet stable without the need for chemical crosslinking agents or other additives that may negatively impact cell function. The scaffold properties can be tuned to some extent by varying factors such as the ECM suspension concentration, mechanical processing methods, or synthesis conditions. In general, the scaffolds are robust and easy to handle, and can be processed as tissues for most standard biological assays, providing a versatile and user-friendly 3D cell culture platform that mimics the native ECM composition. Overall, these straightforward methods for fabricating customized ECM-derived foams and microcarriers may be of interest to both biologists and biomedical engineers as tissue-specific cell-instructive platforms for in vitro and in vivo applications.

The tissue-specific extracellular matrix (ECM) is a key mediator of cell function. This article describes methods for synthesizing pure ECM-derived foams and microcarriers that are stable in culture without the need for chemical crosslinking for applications in advanced 3D in vitro cell culture models or as pro-regenerative bioscaffolds.

Cell function is mediated by interactions with the extracellular matrix (ECM), which has complex tissue-specific composition and architecture. The focus ...

3D Microtissues for Injectable Regenerative Therapy and High-throughput Drug Screening

To upgrade traditional 2D cell culture to 3D cell culture, we have integrated microfabrication with cryogelation technology to produce macroporous microscale cryogels (microcryogels), which can be loaded with a variety of cell types to form 3D microtissues. Herein, we present the protocol to fabricate versatile 3D microtissues and their applications in regenerative therapy and drug screening. Size and shape-controllable microcryogels can be fabricated on an array chip, which can be harvested off-chip as individual cell-loaded carriers for injectable regenerative therapy or be further assembled on-chip into 3D microtissue arrays for high-throughput drug screening. Due to the high elastic nature of these microscale cryogels, the 3D microtissues exhibit great injectability for minimally invasive cell therapy by protecting cells from mechanical shear force during injection. This ensures enhanced cell survival and therapeutic effect in the mouse limb ischemia model. Meanwhile, assembly of 3D microtissue arrays in a standard 384-multi-well format facilitates the use of common laboratory facilities and equipment, enabling high-throughput drug screening on this versatile 3D cell culture platform.

This protocol describes the fabrication of elastic 3D macroporous microcryogels by integrating microfabrication with cryogelation technology. Upon loading with cells, 3D microtissues are generated, which can be readily injected in vivo to facilitate regenerative therapy or assembled into arrays for in vitro high-throughput drug screening.

To upgrade traditional 2D cell culture to 3D cell culture, we have integrated microfabrication with cryogelation technology to produce macroporous ...

Microengineering 3D Collagen Hydrogels with Long-Range Fiber Alignment

Aligned collagen I (COL1) fibers guide tumor cell motility, influence endothelial cell morphology, control stem cell differentiation, and are a hallmark of cardiac and musculoskeletal tissues. To study cell response to aligned microenvironments in vitro, several protocols have been developed to generate COL1 matrices with defined fiber alignment, including magnetic, mechanical, cell-based, and microfluidic methods. Of these, microfluidic approaches offer advanced capabilities such as accurate control over fluid flows and the cellular microenvironment. However, the microfluidic approaches to generate aligned COL1 matrices for advanced in vitro culture platforms have been limited to thin "mats" (&lt;40 &#181;m in thickness) of COL1 fibers that extend over distances less than 500 &#181;m and are not conducive to 3D cell culture applications. Here, we present a protocol to fabricate 3D COL1 matrices (130-250 &#181;m in thickness) with millimeter-scale regions of defined fiber alignment in a microfluidic device. This platform provides advanced cell culture capabilities to model structured tissue microenvironments by providing direct access to the micro-engineered matrix for cell culture.

This protocol demonstrates the use of a microfluidic channel with changing geometry along the fluid flow direction to generate extensional strain (stretching) to align fibers in a 3D collagen hydrogel (&lt;250 &#181;m in thickness). The resulting alignment extends across several millimeters and is influenced by the extensional strain rate.

Aligned collagen I (COL1) fibers guide tumor cell motility, influence endothelial cell morphology, control stem cell differentiation, and are a hallmark ...

Closed System Production of Adherent Cells Based on Dissolvable Microcarriers

Large-Scale Cell Production Based on GMP-Grade Dissolvable Porous Microcarriers

Researchers in the cell and gene therapy (CGT) industry have long faced a formidable challenge in the efficient and large-scale expansion of cells. To address the primary shortcomings of the two-dimensional (2D) planar culturing system, we innovatively developed an automated closed industrial scale cell production (ACISCP) platform based on a GMP-grade, dissolvable, and porous microcarrier for the 3D culture of adherent cells, including human mesenchymal stem/stromal cells (hMSCs), HEK293T cells, and Vero cells. To achieve large-scale expansion, a two-stage expansion was conducted with 5 L and 15 L stirred-tank bioreactors to yield 1.1 x 1010 hMSCs with an overall 128-fold expansion within 9 days. The cells were harvested by completely dissolving the microcarriers, concentrated, washed and formulated with a continuous-flow centrifuge-based cell processing system, and then aliquoted with a cell filling system. Compared with 2D planar culture, there are no significant differences in the quality of hMSCs harvested from 3D culture. We have also applied these dissolvable porous microcarriers to other popular cell types in the CGT sector; specifically, HEK293T cells and Vero cells have been cultivated to peak cell densities of 1.68 x 107 cells/mL and 1.08 x 107 cells/mL, respectively. This study provides a protocol for using a bioprocess engineering platform harnessing the characteristics of GMP-grade dissolvable microcarriers and advanced closed equipment to achieve the industrial-scale manufacturing of adherent cells.

Author Spotlight: Advancing Cell Therapy Manufacturing with Dissolvable Microcarriers 

Here, we present a protocol to achieve the large-scale manufacturing of adherent cells through a fully closed system based on GMP-grade dissolvable microcarriers. The cultivation of human mesenchymal stem cells, HEK293T cells, and Vero cells was validated and met both quantity demands and quality criteria for the cell and gene therapy industry.

Researchers in the cell and gene therapy (CGT) industry have long faced a formidable challenge in the efficient and large-scale expansion of cells. To ...

Evaluating Biofunctional Self-Assembling Peptides for Cell Adhesion and Morphology

Fabrication and Characterization of Colorectal Cancer Organoids from SW1222 Cell Line in Ultrashort Self&#45;Assembling Peptide Matrix

Ultrashort self-assembling peptides (SAPs) can spontaneously form nanofibers that resemble the extracellular matrix. These fibers allow the formation of hydrogels that are biocompatible, biodegradable, and non-immunogenic. We have previously proven that SAPs, when biofunctionalized with protein-derived motifs, can mimic the extracellular matrix characteristics that support colorectal organoid formation. These biofunctional peptide hydrogels retain the original parent peptide's mechanical properties, tunability, and printability while incorporating cues that allow cell-matrix interactions to increase cell adhesion. This paper presents the protocols needed to evaluate and characterize the effects of various biofunctional peptide hydrogels on cell adhesion and lumen formation using an adenocarcinoma cancer cell line able to form colorectal cancer organoids cost-effectively. These protocols will help evaluate biofunctional peptide hydrogel effects on cell adhesion and luminal formation using immunostaining and fluorescence image analysis. The cell line used in this study has been previously utilized for generating organoids in animal-derived matrices.

Author Spotlight: Optimization of Ultrashort Peptide Matrices for Colorectal Cancer Organoids

This protocol aims to evaluate biofunctional self-assembling peptides for cell adhesion, organoid morphology, and gene expression by immunostaining. We will use a colorectal cancer cell line to provide a cost-effective way of obtaining organoids for intensive testing.

Ultrashort self-assembling peptides (SAPs) can spontaneously form nanofibers that resemble the extracellular matrix. These fibers allow the formation of ...

Recombinant Collagen I Peptide Microcarriers for Cell Expansion and Their Potential Use As Cell Delivery System in a Bioreactor Model

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