We are grateful for the excellent technical assistance of Janine Becker, Caroline Bickelmann and Ruth Nickels. This study was funded by a grant of the Deutsche Forschungsgemeinschaft (DFG - German Research Foundation) - LA 2682/7-1.

All procedures were performed according to the National Institute of Health guidelines for the use of experimental animals and followed institutional guidelines (Landesamt f&#252;r Soziales, Gesundheit und Verbraucherschutz, Abt. Lebensmittel- und Veterin&#228;rwesen, Zentralstelle, Saarbr&#252;cken, Germany).
1. Preparation of Surgical Instruments
<ol>
	<li>Keep ready the dissection scissors, surgical forceps, small preparation scissors, fine forceps and a sterile Petri dish with 15 mL Dulbecco&#39;s modified Eagle medium (DMEM; 10% fetal calf serum (FCS), 100 U/mL penicillin, 0.1 mg/mL streptomycin) for harvesting epididymal fat pads.</li>
	<li>Expose the surgical instruments to a disinfecting solution for 5 min. Alternatively, sterilize them (steam sterilization; 121 &#176;C, 20 min).</li>
</ol>
2. Animals and Anesthesia
<ol>
	<li>Choose carefully the strain of the mice as indicated for the study and the subject under investigation. 
	NOTE: In this study we used male wild-type C57BL/6 mice as donors for the harvesting of epididymal fat. Of interest, ad-MVF may be also isolated from transgenic green fluorescent protein (GFP)-positive donor animals (C57BL/6-Tg(CAG-EGFP)1Osb/J)22. This bears the major advantage that the fragments are easily detectable by immunohistochemical staining of GFP after implantation into GFP-negative wild-type mice14.</li>
	<li>Anesthetize the animals with an intraperitoneal injection of xylazine (15 mg/kg) and ketamine (75 mg/kg). Make sure that the animals are deeply anesthetized by performing a toe pinch with no response. Eye lubricant is not indicated as the donor animals are sacrificed after fat harvesting. 
	NOTE: Ensure that analgesia and surgical sterility are in agreement with the respective guidelines of the country and institution where the experiments are planned.</li>
</ol>
3. Harvesting of Epididymal Fat Pads
<ol>
	<li>Transfer the animal to an operation table. Place the animal in a supine position under a surgical stereomicroscope and confirm deep anesthesia using toe pinch.</li>
	<li>Immobilize the paws by taping them to a surgical drape and disinfect the abdomen with disinfecting solution.</li>
	<li>Separate the abdominal skin free of the underlying muscle layer with the dissection scissors.</li>
	<li>Perform a midline laparotomy with the dissection scissors and laterally unfold the flaps of the abdominal wall.</li>
	<li>Bilaterally identify testis, epididymis and the epididymal fat pad using the fine forceps (Figure 1). Do not harm the intestinal structures to prevent fecal contamination of the fat pads.</li>
	<li>Harvest the epididymal fat pads with the small preparation scissors and the fine forceps under the stereomicroscope. Keep a safety margin of several mm between the epididymis and the fat to reduce the risk of accidental epididymal harvesting. 
	NOTE: This procedure can also be performed without a stereomicroscope. However, this may further increase the risk of accidental injury of the epididymis and spermatic cords.</li>
	<li>Transfer the epididymal fat pads into a Petri dish containing 15 mL of DMEM pre-heated at 37 &#176;C for the transport to the cell laboratory.</li>
	<li>Sacrifice the animal by incision of the abdominal aorta or cervical dislocation.</li>
</ol>
4. Isolation of ad-MVF
<ol>
	<li>Prepare three sterile Petri dishes with 15 mL of phosphate-buffered saline (PBS), sterile 14-mL polypropylene (PP) tubes, a sterile 50-mL Erlenmeyer flask, sterile 1.5-mL conical microcentrifuge tubes and sterilized fine scissors.</li>
	<li>Wash the fat pads thrice in Petri dishes with 15 mL of PBS under a laminar flow hood.</li>
	<li>Transfer the fat into a 14-mL PP tube. Determine the volume of harvested fat tissue (in mL) by means of the tube scale. Mince the fat tissue mechanically with the fine scissors until a homogeneous tissue suspension is obtained.</li>
	<li>Transfer the minced tissue with two volumes of collagenase NB4G (0.5 U/mL PBS) into a 50-mL Erlenmeyer flask by means of 10-mL measuring pipette. The total volume becomes thrice the volume of fat tissue measured in step 4.3. Perform tissue digestion in an incubator for 10 min under vigorous stirring by means of an automated stirrer (size of magnetic stir bar: 25 mm) at 37 &#176;C and humidified atmospheric conditions with 5% CO2.</li>
	<li>Observe a small fraction (10 &#181;L) of the digested tissue under a microscope to judge whether the digestion can be stopped. Ascertain that the digestate mainly contains &#34;free&#34; ad-MVF next to single cells, indicating the appropriate point to stop the enzymatic digestion process (Figure 2). 
	NOTE: This step requires experience with the procedure and is critical for the quality of isolated ad-MVF. Prolonged fat digestion results in a single-cell suspension without ad-MVF.</li>
	<li>Neutralize the enzyme with two volumes of PBS/20% FCS. The total volume becomes thrice the volume of the cell-vessel suspension in step 4.4. Transfer the cell-vessel suspension back into new PP tubes.</li>
	<li>Incubate the suspension for 5 min at 37 &#176;C to separate ad-MVF from remaining fat by gravity. Then carefully remove the main fat supernatant with a 1-mL precision pipette. Repeat this cycle several times with a 100-&#181;L precision pipette until the suspension appears to be fat-free.</li>
	<li>Put a 500-&#181;m filter on top of a 50-mL conical centrifuge tube. Transfer the cell-vessel suspension with a 10-mL measuring pipette from the 14-mL PP tubes onto the filter membrane to remove remaining fat clots. Transfer the filtered suspension into new 14-mL PP tubes according to the number of individual ad-MVF isolates for the planned experiments. 
	NOTE: For in vitro assays focusing on microvascular network formation, it may be beneficial to further purify the cell-vessel suspension to improve the imaging quality during microscopic analyses. For this purpose, the suspension may be additionally filtered once with a 20-&#181;m filter to remove single cells from the ad-MVF collected on the filter.</li>
	<li>Centrifuge the cell-vessel suspension (600 x g, 5 min, room temperature) to obtain a pellet containing ad-MVF.</li>
	<li>After centrifugation, remove the supernatant until 1 mL is left. Resuspend the pellet with ad-MVF in this 1 mL and transfer the suspension into a 1.5-mL conical microcentrifuge tube.</li>
	<li>Centrifuge the ad-MVF suspension in the microcentrifuge tube (600 x g, 5 min) to obtain a pellet.</li>
	<li>Remove the supernatant to resuspend the pellet in the required final volume of PBS / 20% FCS. 
	NOTE: The number of ad-MVF per isolate can be assessed by microscopic counting. For this purpose, 1/10 of the final cell-vessel suspension is diluted 1:10 in PBS and 100 &#181;L of this dilution are transferred into a well of a 96-well plate. The entire number of ad-MVF exhibiting a vessel-like morphology is then counted and extrapolated to the whole isolate. The required ad-MVF concentration and purification can be individually adapted to the respective in vitro or in vivo application of the ad-MVF. This can include the embedding of ad-MVF in collagen gels for the in vitro analysis of microvascular network formation or the seeding of ad-MVF on scaffolds for in vivo implantation into tissue defects.</li>
</ol>

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The authors declare that they have no competing financial interests.

In this study we present a well-established protocol for the isolation of ad-MVF. Obtaining ad-MVF from murine adipose tissue is a straightforward procedure with a few critical steps. Mice exhibit different subcutaneous and intraabdominal fat deposits. As previously described for rats, the most suitable fat source for the isolation of ad-MVF are the epididymal fat pads due to their size, homogeneous structure and minimal contamination with larger blood vessels11,12. In contrast, subcutaneous fat deposits in mice are much smaller and more adherent to the surrounding tissue. However, the excision of epididymal fat pads bears the risk of accidental injury of the epididymis and spermatic cords, resulting in contamination of the harvested tissue with sperms. Therefore, the identification of epididymis, testis and spermatic cords is of major importance in this procedure. Moreover, the fat pads should be rapidly processed after harvesting to minimize ex vivo tissue damage. Finally, it should be considered that there are major age-dependent variations in the vascularization of murine epididymal fat pads23. In our experience, the use of 6- to 12-month-old male donor mice provides sufficient amounts of ad-MVF for in vitro and in vivo studies. However, it may also be possible to isolate ad-MVF from younger mice, which exhibit smaller but highly vascularized fat pads24.
The enzymatic digestion of the harvested epididymal fat pads crucially determines the final quality of isolated ad-MVF. Too short exposition to collagenase leads to incomplete separation of ad-MVF from surrounding adipocytes. On the other hand, prolonged digestion results in the destruction of ad-MVF to a single cell suspension, the so-called stromal vascular fraction (SVF). The SVF is a heterogeneous cell population containing mesenchymal stem cells, endothelial cells, pericytes, macrophages and many other cell types with regenerative potential through paracrine and differentiation mechanisms25. The use of ad-MVF exhibits considerable advantages when compared to SVF-based approaches. The fully functional microvessel morphology of ad-MVF allows their immediate implantation for in vivo experiments without time-consuming and complex in vitro cultivation. Moreover, the isolation of the SVF usually requires the digestion of the tissue with collagenase up to 90 min26. In contrast, tissue degradation should not exceed 10 min for the isolation of ad-MVF. This supports the idea of intraoperative one-step applications of ad-MVF9. For this purpose, ad-MVF may be automatically isolated from autologous lipoaspirates and transferred back into a tissue defect of the patient without leaving the operation theatre. This is a realistic goal considering the fact that the automated isolation of SVF from lipoaspirates is already a clinically established procedure in plastic surgery27,28,29.
Recently, we have demonstrated a strong effect of ad-MVF on the in vivo vascularization of bone14 and skin15 substitutes. Of interest, we found that it is beneficial not to remove single cells from the ad-MVF isolate, because they may provide a more physiological environment that contributes to the high vascularization capacity of ad-MVF13. Therefore, we only used a 500-&#181;m filter during the isolation of ad-MVF for the removal of larger non-digested fat clots. Moreover, we found that ad-MVF induce lymphangiogenesis in implanted skin substitutes15. Hence, ad-MVF may also be promising building blocks for lymphatic tissue engineering and lymphedema therapy.
For the successful translation of these experimental findings into clinical practice, the next logical step is to clarify whether human adipose tissue is equally suited for the isolation of ad-MVF as rodent fat. In this context, a critical issue is the amount of fat required to isolate large amounts of ad-MVF. The herein described protocol resulted in approximately 40,000 ad-MVF per mL fat tissue. Accordingly, 3 mL fat tissue would be necessary to prevascularize 1 cm2 of dermal skin substitute15. Hence, this approach may not be feasible for the treatment of larger wounds or severely ill patients due to a limited amount of available fat tissue for ad-MVF isolation.
Beside a broad spectrum of potential in vivo applications in tissue engineering and regenerative medicine, ad-MVF are also suitable for in vitro modeling of angiogenic processes. For instance, rat ad-MVF have been cultured in three-dimensional collagen hydrogels to study the formation of new microvascular networks30. Similar approaches have been used to evaluate sophisticated imaging techniques for the visualization of neovessel growth13 or interstitial matrix remodeling during angiogenesis31.
Taken together, these findings indicate that ad-MVF are not only promising vascularization and lymphangiogenic units for future therapeutic strategies, but also for basic research in the field of angiogenesis and vascular physiology.

Tissue engineering focuses on the fabrication of tissue and organ substitutes that maintain, restore or augment the function of inoperable in vivo counterparts1,2. The fate of engineered tissue constructs crucially depends on an adequate vascularization3. Microvascular networks within these constructs should be hierarchically organized with arterioles, capillaries, and venules to allow efficient blood perfusion after inosculation to the recipient&#39;s vasculature4. The generation of such networks is among the key challenges in tissue engineering. For this purpose, a broad spectrum of experimental vascularization strategies has been introduced over the last two decades5,6.
Angiogenic approaches stimulate the ingrowth of recipient microvessels into engineered tissues by means of structural or physicochemical scaffold modification, such as the incorporation of growth factors7. However, for the vascularization of large three-dimensional constructs, angiogenesis-dependent strategies are markedly limited by slow growth rates of developing microvessels8.
In contrast, the concept of prevascularization aims for the generation of functional microvascular networks within tissue constructs prior to their implantation9. Conventional prevascularization involves the co-culture of vessel-forming cells, such as endothelial cells, mural cells or stem cells10, within scaffolds. After microvascular network formation, the prevascularized constructs can then be implanted into tissue defects. Noteworthy, this prevascularization approach is difficult to apply in a clinical setting, because it is based on complex and time-consuming in vitro procedures, which are restricted by major regulatory hurdles9. Accordingly, there is still a need for the development of novel prevascularization strategies that are more suitable for a broad clinical application.
Such a prevascularization strategy may be the application of adipose tissue-derived microvascular fragments (ad-MVF). ad-MVF represent potent vascularization units that can be harvested in large amounts from the fat tissue of rats11,12 and mice13. They consist of arteriolar, capillary, and venular vessel segments, which exhibit a physiological microvessel morphology with a lumen and stabilizing perivascular cells14,15. This unique feature allows the immediate implantation of ad-MVF-seeded scaffolds into tissue defects without precultivation. There, the ad-MVF rapidly reassemble into functional microvascular networks. Furthermore, ad-MVF represent a rich source of mesenchymal stem cells16, which may additionally contribute to their striking regenerative capacity. Accordingly, ad-MVF are increasingly used in different fields of tissue engineering14,15,17,18,19,20,21.
The isolation of ad-MVF has originally been established in rats11,12. Herein, we describe a protocol, which allows the standardized isolation of murine ad-MVF from epididymal fat pads. This may provide further insights into molecular mechanisms underlying ad-MVF function by using transgenic mouse models.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.5-mL conical microcentrifuge tube</td>
			<td>VWR, Kelsterbach, Germany</td>
			<td>700-5239</td>
			<td></td>
		</tr>
		<tr>
			<td>100-&#181;L precision pipette</td>
			<td>Eppendorf, Hamburg, Germany</td>
			<td>4920000059</td>
			<td></td>
		</tr>
		<tr>
			<td>10-mL measuring pipette</td>
			<td>Costar, Corning Inc., New York, USA</td>
			<td>4488</td>
			<td></td>
		</tr>
		<tr>
			<td>14-mL PP tubes</td>
			<td>Greiner bio-one, Frickenhausen, Germany</td>
			<td>187261</td>
			<td></td>
		</tr>
		<tr>
			<td>1-mL precision pipette</td>
			<td>Eppendorf, Hamburg, Germany</td>
			<td>4920000083</td>
			<td></td>
		</tr>
		<tr>
			<td>500-&#181;m filter (pluriStrainer 500 &#181;m)</td>
			<td>HISS Diagnostics, Freiburg, Germany</td>
			<td>43-50500-03</td>
			<td></td>
		</tr>
		<tr>
			<td>50-mL conical centrifuge tube</td>
			<td>Greiner bio-one, Frickenhausen, Germany</td>
			<td>227261</td>
			<td></td>
		</tr>
		<tr>
			<td>50-mL Erlenmeyer flask</td>
			<td>VWR, Kelsterbach, Germany</td>
			<td>214-0211</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well plate</td>
			<td>Greiner bio-one, Frickenhausen, Germany</td>
			<td>65518</td>
			<td></td>
		</tr>
		<tr>
			<td>cell detachment solution (Accutase)</td>
			<td>eBioscience, San Diego, CA USA</td>
			<td>00-4555-56</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6 mice</td>
			<td>Charles River, Cologne, Germany</td>
			<td>027</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6-Tg(CAG-EGFP)1Osb/J mice</td>
			<td>The Jackson Laboratory, Bar Harbor, USA</td>
			<td>003291</td>
			<td></td>
		</tr>
		<tr>
			<td>CD117-FITC</td>
			<td>BD Biosciences, Heidelberg, Germany</td>
			<td>553373</td>
			<td></td>
		</tr>
		<tr>
			<td>CD31-PE</td>
			<td>BD Biosciences, Heidelberg, Germany</td>
			<td>553354</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase NB4G&#160;</td>
			<td>Serva Electrophoresis GmbH, Heidelberg, Germany</td>
			<td>17465.02</td>
			<td>Lot tested by manufacturer for enzymatic activity</td>
		</tr>
		<tr>
			<td>Dissection scissors</td>
			<td>Braun Aesculap AG &#38;CoKG, Melsungen, Germany</td>
			<td>BC 601</td>
			<td></td>
		</tr>
		<tr>
			<td>DNA-binding dye (Bisbenzimide H33342)</td>
			<td>Sigma-Aldrich, Taufkirchen, Germany</td>
			<td>B2261</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s modified Eagle medium (DMEM)&#160;</td>
			<td>PAN Biotech, Rickenbach, Germany</td>
			<td>P04-03600</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal calf serum (FCS)</td>
			<td>Biochrom GmbH, Berlin, Germany</td>
			<td>S0615</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>S&#38;T AG, Neuhausen, Switzerland</td>
			<td>FRS-15 RM-8</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine scissors</td>
			<td>World Precision Instrumets, Sarasota, FL, USA</td>
			<td>503261</td>
			<td></td>
		</tr>
		<tr>
			<td>Dermal skin substitute (Integra)</td>
			<td>Integra Life Sciences, Sain Priest, France</td>
			<td>62021</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine&#160;</td>
			<td>Serumwerk Bernburg AG, Bernburg, Germany</td>
			<td>7005294</td>
			<td></td>
		</tr>
		<tr>
			<td>M-IgG2akAL488&#160;&#160;</td>
			<td>eBioscience, San Diego, CA USA</td>
			<td>53-4724-80</td>
			<td></td>
		</tr>
		<tr>
			<td>Octeniderm (disinfecting solution)</td>
			<td>Sch&#252;lke &#38; Mayer, Norderstedt, Germany</td>
			<td>118211</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin/Streptomycin</td>
			<td>Biochrom, Berlin, Germany</td>
			<td>A2213</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Greiner bio-one, Frickenhausen, Germany</td>
			<td>664160</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate-buffered saline (PBS)</td>
			<td>Lonza Group, Basel, Switzerland</td>
			<td>17-516F</td>
			<td></td>
		</tr>
		<tr>
			<td>pluriStrainer 20-&#181;m (20 &#181;m filter)</td>
			<td>HISS Diagnostics, Freiburg, Germany</td>
			<td>43-50020-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat-IgG2akFITC</td>
			<td>BD Biosciences, Heidelberg, Germany</td>
			<td>553988</td>
			<td></td>
		</tr>
		<tr>
			<td>Rat-IgG2akPE</td>
			<td>BD Biosciences, Heidelberg, Germany</td>
			<td>553930</td>
			<td></td>
		</tr>
		<tr>
			<td>Small preparation scissors</td>
			<td>S&#38;T AG, Neuhausen, Switzerland</td>
			<td>SDC-15 R-8S</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical forceps</td>
			<td>Braun Aesculap AG &#38;CoKG, Melsungen, Germany</td>
			<td>BD510R</td>
			<td></td>
		</tr>
		<tr>
			<td>Tape (Heftpflaster Seide) 1.25 cm</td>
			<td>Fink &#38; Walter GmbH, Mechweiler, Germany</td>
			<td>1671801</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine&#160;</td>
			<td>Bayer Vital GmbH, Leverkusen, Germany</td>
			<td>1320422</td>
			<td></td>
		</tr>
		<tr>
			<td>&#945;-SMA-AL488</td>
			<td>eBioscience, San Diego, CA USA</td>
			<td>53-9760-82</td>
			<td>Intracellular labeling additionally requires Cytofix/Cytoperm (BD Biosciences, Heidelberg, Germany; #554722)</td>
		</tr>
	</tbody>
</table>

isolation of murine adipose tissue-derived microvascular fragments as vascularization units for tissue engineering

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

In the present study we performed six ad-MVF isolation procedures with fat tissue from 7- to 12-month-old male wild-type C57BL/6 mice (mean body weight: 35 &#177; 1 g). Figure 1 illustrates the harvesting of murine epididymal fat pads with subsequent mechanical and enzymatic ad-MVF isolation. The time required for the harvesting of fat was 30 min and for the isolation of ad-MVF was 120 min. In total, the procedure took 150 min.
We harvested 1.2 &#177; 0.1 mL of adipose tissue per donor animal. This amount of fat allowed the isolation of 42,000 &#177; 2,000 ad-MVF per mL. The mean length of the isolated ad-MVF was 42 &#177; 1 &#181;m (Figure 2A). The duration of enzymatic digestion was determined by means of microscopic control as shown in Figure 2B. ad-MVF exhibited a typical microvessel morphology with hierarchical vessel segments (Figure 2C).
We additionally characterized the ad-MVF by means of flow cytometry. For this purpose, ad-MVF were further digested in cell detachment solution for 30 min into single cells15. The flow cytometric analyses revealed that ad-MVF contain 26 &#177; 2% CD31-positive endothelial cells, 17 &#177; 2% &#945;-smooth muscle actin (SMA)-positive perivascular cells, and 9 &#177; 1% cells positive for the mesenchymal stem cell marker CD117 (Figure 3A-C). After seeding on a dermal skin substitute, which was fixed and sectioned immediately for histological analyses, larger ad-MVF were mainly localized on the implant&#39;s surface (Figure 3D). However, capillary vessel segments could also be detected within its pores. Immunohistochemical staining of seeded ad-MVF further revealed physiological microvessel configuration with arteriolar, venular and capillary-like fragments (Figure 3E).
<img alt="Figure 1" src="/files/ftp_upload/55721/55721fig1.jpg" /> 
Figure 1: ad-MVF Isolation. A) Following midline laparotomy, the epididymal fat pads (asterisks) are identified. The adipose tissue is harvested with a small preparation scissors and the animal is sacrificed by incision of the abdominal aorta (arrowhead). B) Mechanical mincing of the fat pads with fine scissors until the tissue suspension appears homogeneous. C) For enzymatic digestion, collagenase is added to the tissue suspension. D) The cell-vessel suspension is incubated and the supernatant fat layer is discarded. E) After centrifugation, the pellet containing ad-MVF is resuspended and can be used for different applications, such as the in vitro analysis of microvascular network formation or the seeding of ad-MVF on scaffolds for in vivo implantation into tissue defects (F). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55721/55721fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55721/55721fig2.jpg" /> 
Figure 2: Morphological Characteristics of ad-MVF. A) Length distribution of freshly isolated ad-MVF. Mean &#177; standard error of the mean (n = 6). B) Microscopic image of a cell-vessel suspension smear with large (arrow), medium-sized (double arrows) and small (black arrowheads) ad-MVF as well as single cells (white arrowheads). Scale bar = 110 &#181;m. Higher magnification reveals that ad-MVF exhibit a mature microvessel morphology with hierarchical microvessel segments (C, inset of B). Scale bar: C = 50 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55721/55721fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/55721/55721fig3.jpg" /> 
Figure 3: Flow Cytometric, Histological and Immunohistochemical Characteristics of ad-MVF. A-C) Flow cytometric analyses of freshly isolated ad-MVF illustrating CD31-positive endothelial cell (A), &#945;-SMA-positive perivascular cell (B) and CD117-positive mesenchymal stem cell (C) fractions. Appropriate isotype-identical controls were used to adjust threshold levels. Mean &#177; standard error of the mean (n = 6). D) Hematoxylin and eosin-stained section of a seeded dermal skin substitute. A few capillary vessel segments are located in the implant&#39;s pores (arrowheads). Larger microvessels with venular (double arrow) or arteriolar (arrow) morphology are localized on its surface. Scale bar = 40 &#181;m. Broken line = implant border. E) Characterization of ad-MVF by means of immunohistochemical detection of the endothelial cell marker CD31 (red) and the perivascular cell marker &#945;-SMA (green). Cell nuclei were stained with a DNA-binding fluorescent dye (blue). Arteriolar ad-MVF (arrow) are characterized by a thicker &#945;-SMA-positive cell layer when compared to venular fragments with a larger lumen (double arrow). Arrowheads = capillary ad-MVF. Scale bar = 25 &#181;m. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55721/55721fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Isolation of Murine Adipose Tissue-derived Microvascular Fragments as Vascularization Units for Tissue Engineering at JoVE.com

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

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

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

isolation-murine-adipose-tissue-derived-microvascular-fragments-as

Nanyang Technological University

Research

JoVE Journal

Bioengineering

12.3K Views.  Saarland University. We present a protocol to isolate adipose tissue-derived microvascular fragments that represent promising vascularization units. They can be rapidly isolated, do not require in vitro processing and, thus, may be used for one-step prevascularization in different fields of tissue engineering.

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

Fabrication of Biologically Derived Injectable Materials for Myocardial Tissue Engineering

This protocol provides methods for the preparation of an injectable extracellular matrix (ECM) gel for myocardial tissue engineering applications. Briefly, decellularized tissue is lyophilized, milled, enzymatically digested, and then brought to physiological pH. The lyophilization removes all water content from the tissue, resulting in dry ECM that can be ground into a fine powder with a small mill. After milling, the ECM powder is digested with pepsin to form an injectable matrix. After adjustment to pH 7.4, the liquid matrix material can be injected into the myocardium. Results of previous characterization assays have shown that matrix gels produced from decellularized pericardial and myocardial tissue retain native ECM components, including diverse proteins, peptides and glycosaminoglycans. Given the use of this material for tissue engineering, in vivo characterization is especially useful; here, a method for performing an intramural injection into the left ventricular (LV) free wall is presented as a means of analyzing the host response to the matrix gel in a small animal model. Access to the chest cavity is gained through the diaphragm and the injection is made slightly above the apex in the LV free wall. The biologically derived scaffold can be visualized by biotin-labeling before injection and then staining tissue sections with a horse radish peroxidase-conjugated neutravidin and visualizing via diaminobenzidine (DAB) staining. Analysis of the injection region can also be done with histological and immunohistochemical staining. In this way, the previously examined pericardial and myocardial matrix gels were shown to form fibrous, porous networks and promote vessel formation within the injection region.

Methods for preparing an injectable matrix gel from decellularized tissue and injecting it into rat myocardium in vivo are described.

This protocol provides methods for the preparation of an injectable extracellular matrix (ECM) gel for myocardial tissue engineering applications.  ...

Postproduction Processing of Electrospun Fibres for Tissue Engineering

Electrospinning is a commonly used and versatile method to produce scaffolds (often biodegradable) for 3D tissue engineering.1, 2, 3 Many tissues in vivo undergo biaxial distension to varying extents such as skin, bladder, pelvic floor and even the hard palate as children grow. In producing scaffolds for these purposes there is a need to develop scaffolds of appropriate biomechanical properties (whether achieved without or with cells) and which are sterile for clinical use. The focus of this paper is not how to establish basic electrospinning parameters (as there is extensive literature on electrospinning) but on how to modify spun scaffolds post production to make them fit for tissue engineering purposes - here thickness, mechanical properties and sterilisation (required for clinical use) are considered and we also describe how cells can be cultured on scaffolds and subjected to biaxial strain to condition them for specific applications.
Electrospinning tends to produce thin sheets; as the electrospinning collector becomes coated with insulating fibres it becomes a poor conductor such that fibres no longer deposit on it. Hence we describe approaches to produce thicker structures by heat or vapour annealing increasing the strength of scaffolds but not necessarily the elasticity. Sequential spinning of scaffolds of different polymers to achieve complex scaffolds is also described. Sterilisation methodologies can adversely affect strength and elasticity of scaffolds. We compare three methods for their effects on the biomechanical properties on electrospun scaffolds of poly lactic-co-glycolic acid (PLGA).
Imaging of cells on scaffolds and assessment of production of extracellular matrix (ECM) proteins by cells on scaffolds is described. Culturing cells on scaffolds in vitro can improve scaffold strength and elasticity but the tissue engineering literature shows that cells often fail to produce appropriate ECM when cultured under static conditions. There are few commercial systems available that allow one to culture cells on scaffolds under dynamic conditioning regimes - one example is the Bose Electroforce 3100 which can be used to exert a conditioning programme on cells in scaffolds held using mechanical grips within a media filled chamber.4 An approach to a budget cell culture bioreactor for controlled distortion in 2 dimensions is described. We show that cells can be induced to produce elastin under these conditions. Finally assessment of the biomechanical properties of processed scaffolds cultured with or without cells is described.

Electrospun scaffolds can be processed post production for tissue engineering applications. Here we describe methods for spinning complex scaffolds (by consecutive spinning), for making thicker scaffolds (by multi-layering using heat or vapour annealing), for achieving sterility (aseptic production or sterilisation post production) and for achieving appropriate biomechanical properties.

Electrospinning is a commonly used and versatile method to produce  scaffolds (often biodegradable) for 3D tissue engineering.1, 2, 3 Many tissues in vivo ...

Engineering Skeletal Muscle Tissues from Murine Myoblast Progenitor Cells and Application of Electrical Stimulation

Engineered muscle tissues can be used for several different purposes, which include the production of tissues for use as a disease model in vitro, e.g. to study pressure ulcers, for regenerative medicine and as a meat alternative 1. The first reported 3D muscle constructs have been made many years ago and pioneers in the field are Vandenburgh and colleagues 2,3. Advances made in muscle tissue engineering are not only the result from the vast gain in knowledge of biochemical factors, stem cells and progenitor cells, but are in particular based on insights gained by researchers that physical factors play essential roles in the control of cell behavior and tissue development. State-of-the-art engineered muscle constructs currently consist of cell-populated hydrogel constructs. In our lab these generally consist of murine myoblast progenitor cells, isolated from murine hind limb muscles or a murine myoblast cell line C2C12, mixed with a mixture of collagen/Matrigel and plated between two anchoring points, mimicking the muscle ligaments. Other cells may be considered as well, e.g. alternative cell lines such as L6 rat myoblasts 4, neonatal muscle derived progenitor cells 5, cells derived from adult muscle tissues from other species such as human 6 or even induced pluripotent stem cells (iPS cells) 7. Cell contractility causes alignment of the cells along the long axis of the construct 8,9 and differentiation of the muscle progenitor cells after approximately one week of culture. Moreover, the application of electrical stimulation can enhance the process of differentiation to some extent 8. Because of its limited size (8 x 2 x 0.5 mm) the complete tissue can be analyzed using confocal microscopy to monitor e.g. viability, differentiation and cell alignment. Depending on the specific application the requirements for the engineered muscle tissue will vary; e.g. use for regenerative medicine requires the up scaling of tissue size and vascularization, while to serve as a meat alternative translation to other species is necessary.

Engineered muscle tissue has great potential in regenerative medicine, as disease model and also as an alternative source for meat. Here we describe the engineering of a muscle construct, in this case from mouse myoblast progenitor cells, and the stimulation by electrical pulses.

Engineered muscle tissues can be used for several different purposes, which include the production of tissues for use as a disease model in vitro, e.g. to ...

Tissue Engineering of the Intestine in a Murine Model

Tissue-engineered small intestine (TESI) has successfully been used to rescue Lewis rats after massive small bowel resection, resulting in return to preoperative weights within 40 days.1 In humans, massive small bowel resection can result in short bowel syndrome, a functional malabsorptive state that confers significant morbidity, mortality, and healthcare costs including parenteral nutrition dependence, liver failure and cirrhosis, and the need for multivisceral organ transplantation.2 In this paper, we describe and document our protocol for creating tissue-engineered intestine in a mouse model with a multicellular organoid units-on-scaffold approach. Organoid units are multicellular aggregates derived from the intestine that contain both mucosal and mesenchymal elements,3 the relationship between which preserves the intestinal stem cell niche.4 In ongoing and future research, the transition of our technique into the mouse will allow for investigation of the processes involved during TESI formation by utilizing the transgenic tools available in this species.5The availability of immunocompromised mouse strains will also permit us to apply the technique to human intestinal tissue and optimize the formation of human TESI as a mouse xenograft before its transition into humans. Our method employs good manufacturing practice (GMP) reagents and materials that have already been approved for use in human patients, and therefore offers a significant advantage over approaches that rely upon decellularized animal tissues. The ultimate goal of this method is its translation to humans as a regenerative medicine therapeutic strategy for short bowel syndrome.

This article and the accompanying video present our protocol for generating tissue-engineered intestine in the mouse, using an organoid units-on-scaffold approach.

Tissue-engineered small intestine (TESI) has successfully been used to rescue Lewis rats after massive small bowel resection, resulting in return to ...

Capillary Force Lithography for Cardiac Tissue Engineering

Cardiovascular disease remains the leading cause of death worldwide1. Cardiac tissue engineering holds much promise to deliver groundbreaking medical discoveries with the aims of developing functional tissues for cardiac regeneration as well as in vitro screening assays. However, the ability to create high-fidelity models of heart tissue has proven difficult. The heart&#8217;s extracellular matrix (ECM) is a complex structure consisting of both biochemical and biomechanical signals ranging from the micro- to the nanometer scale2. Local mechanical loading conditions and cell-ECM interactions have recently been recognized as vital components in cardiac tissue engineering3-5.
A large portion of the cardiac ECM is composed of aligned collagen fibers with nano-scale diameters that significantly influences tissue architecture and electromechanical coupling2. Unfortunately, few methods have been able to mimic the organization of ECM fibers down to the nanometer scale. Recent advancements in nanofabrication techniques, however, have enabled the design and fabrication of scalable scaffolds that mimic the in vivo structural and substrate stiffness cues of the ECM in the heart6-9.
Here we present the development of two reproducible, cost-effective, and scalable nanopatterning processes for the functional alignment of cardiac cells using the biocompatible polymer poly(lactide-co-glycolide) (PLGA)8 and a polyurethane (PU) based polymer. These anisotropically nanofabricated substrata (ANFS) mimic the underlying ECM of well-organized, aligned tissues and can be used to investigate the role of nanotopography on cell morphology and function10-14.
Using a nanopatterned (NP) silicon master as a template, a polyurethane acrylate (PUA) mold is fabricated. This PUA mold is then used to pattern the PU or PLGA hydrogel via UV-assisted or solvent-mediated capillary force lithography (CFL), respectively15,16. Briefly, PU or PLGA pre-polymer is drop dispensed onto a glass coverslip and the PUA mold is placed on top. For UV-assisted CFL, the PU is then exposed to UV radiation (&#955; = 250-400 nm) for curing. For solvent-mediated CFL, the PLGA is embossed using heat (120 &#176;C) and pressure (100 kPa). After curing, the PUA mold is peeled off, leaving behind an ANFS for cell culture. Primary cells, such as neonatal rat ventricular myocytes, as well as human pluripotent stem cell-derived cardiomyocytes, can be maintained on the ANFS2.

In this protocol, we demonstrate the fabrication of biomimetic cardiac cell culture substrata made from two distinct polymeric materials using capillary force lithography. The described methods provide a scalable, cost-effective technique to engineer the structure and function of macroscopic cardiac tissues for in vitro and in vivo applications.

Cardiovascular disease remains the leading cause of death worldwide1. Cardiac tissue engineering holds much promise to deliver groundbreaking medical ...

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. ...

Preparation of Thermoresponsive Nanostructured Surfaces for Tissue Engineering

Thermoresponsive poly(N-isopropylacrylamide) (PIPAAm)-immobilized surfaces for controlling cell adhesion and detachment were fabricated by the Langmuir-Schaefer method. Amphiphilic block copolymers composed of polystyrene and PIPAAm (St-IPAAms) were synthesized by reversible addition-fragmentation chain transfer (RAFT) radical polymerization. A chloroform solution of St-IPAAm molecules was gently dropped into a Langmuir-trough apparatus, and both barriers of the apparatus were moved horizontally to compress the film to regulate its density. Then, the St-IPAAm Langmuir film was horizontally transferred onto a hydrophobically modified glass substrate by a surface-fixed device. Atomic force microscopy images clearly revealed nanoscale sea-island structures on the surface. The strength, rate, and quality of cell adhesion and detachment on the prepared surface were modulated by changes in temperature across the lower critical solution temperature range of PIPAAm molecules. In addition, a two-dimensional cell structure (cell sheet) was successfully recovered on the optimized surfaces. These unique PIPAAm surfaces may be useful for controlling the strength of cell adhesion and detachment.

Nanoscaled sea-island surfaces composed of thermoresponsive block copolymers were fabricated by the Langmuir-Schaefer method for controlling spontaneous cell adhesion and detachment. Both the preparation of the surface and the adhesion and detachment of cells on the surface were visualized.

Thermoresponsive poly(N-isopropylacrylamide) (PIPAAm)-immobilized surfaces for controlling cell adhesion and detachment were fabricated by the ...

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and trade one defect for another. Tissue engineering holds the promise to address this increasing demand. However, most tissue engineering strategies fail to generate stable and functional tissue substitutes because of poor vascularization. This paper focuses on an in vivo tissue engineering chamber model of intrinsic vascularization where a perfused artery and a vein either as an arteriovenous loop or a flow-through pedicle configuration is directed inside a protected hollow chamber. In this chamber-based system angiogenic sprouting occurs from the arteriovenous vessels and this system attracts ischemic and inflammatory driven endogenous cell migration which gradually fills the chamber space with fibro-vascular tissue. Exogenous cell/matrix implantation at the time of chamber construction enhances cell survival and determines specificity of the engineered tissues which develop. Our studies have shown that this chamber model can successfully generate different tissues such as fat, cardiac muscle, liver and others. However, modifications and refinements are required to ensure target tissue formation is consistent and reproducible. This article describes a standardized protocol for the fabrication of two different vascularized tissue engineering chamber models in vivo.

Tissue Engineering by Intrinsic Vascularization in an In Vivo Tissue Engineering Chamber

This is a guideline for constructing in vivo vascularized tissue using a microsurgical arteriovenous loop or a flow-through pedicle configuration inside a tissue engineering chamber. The vascularized tissues generated can be employed for organ regeneration and replacement of tissue defects, as well as for drug testing and disease modeling.

In reconstructive surgery, there is a clinical need for an alternative to the current methods of autologous reconstruction which are complex, costly and ...

A Microfluidic Platform for High-throughput Single-cell Isolation and Culture

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet high-throughput, method to perform single-cell culture experiments. Here, we report a microfluidic chip-based strategy for high-efficiency single-cell isolation (~77%) and demonstrate its capability of performing long-term single-cell culture (up to 7 d) and cellular heterogeneity analysis using clonogenic assay. These applications were demonstrated with KT98 mouse neural stem cells, and A549 and MDA-MB-435 human cancer cells. High single-cell isolation efficiency and long-term culture capability are achieved by using different sizes of microwells on the top and bottom of the microfluidic channel. The small microwell array is designed for precisely isolating single-cells, and the large microwell array is used for single-cell clonal culture in the microfluidic chip. This microfluidic platform constitutes an attractive approach for single-cell culture applications, due to its flexibility of adjustable cell culture spaces for different culture strategies, without decreasing isolation efficiency.

Here, we present a protocol for isolating and culturing single cells with a microfluidic platform, which utilizes a new microwell design concept to allow for high-efficiency single cell isolation and long-term clonal culture.

Studying the heterogeneity of single cells is crucial for many biological questions, but is technically difficult. Thus, there is a need for a simple, yet ...

Two Methods for Decellularization of Plant Tissues for Tissue Engineering Applications

The autologous, synthetic, and animal-derived grafts currently used as scaffolds for tissue replacement have limitations due to low availability, poor biocompatibility, and cost. Plant tissues have favorable characteristics that make them uniquely suited for use as scaffolds, such as high surface area, excellent water transport and retention, interconnected porosity, preexisting vascular networks, and a wide range of mechanical properties. Two successful methods of plant decellularization for tissue engineering applications are described here. The first method is based on detergent baths to remove cellular matter, which is similar to previously established methods used to clear mammalian tissues. The second is a detergent-free method adapted from a protocol that isolates leaf vasculature and involves the use of a heated bleach and salt bath to clear the leaves and stems. Both methods yield scaffolds with comparable mechanical properties and low cellular metabolic impact, thus allowing the user to select the protocol which better suits their intended application.

Here we present, and contrast two protocols used to decellularize plant tissues: a detergent-based approach and a detergent-free approach. Both methods leave behind the extracellular matrix of the plant tissues used, which can then be utilized as scaffolds for tissue engineering applications.

The autologous, synthetic, and animal-derived grafts currently used as scaffolds for tissue replacement have limitations due to low availability, poor ...