Name: isolation of murine adipose tissue-derived microvascular fragments as vascularization units for tissue engineering
Uploaded: 2017-04-30T13:00:00.000+00:00
Duration: 7 min 9 s
Description: Watch this Scientific Journal Video about Isolation of Murine Adipose Tissue-derived Microvascular Fragments as Vascularization Units for Tissue Engineering at JoVE.com

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><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-&amp;micro;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-&amp;micro;m filter (pluriStrainer 500 &amp;micro;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&amp;nbsp;</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 &amp;amp;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&#039;s modified Eagle medium (DMEM)&amp;nbsp;</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&amp;amp;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&amp;nbsp;</td><td>Serumwerk Bernburg AG, Bernburg, Germany</td><td>7005294</td><td></td></tr><tr><td>M-IgG2akAL488&amp;nbsp;&amp;nbsp;</td><td>eBioscience, San Diego, CA USA</td><td>53-4724-80</td><td></td></tr><tr><td>Octeniderm (disinfecting solution)</td><td>Sch&amp;uuml;lke &amp;amp; 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-&amp;micro;m (20 &amp;micro;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&amp;amp;T AG, Neuhausen, Switzerland</td><td>SDC-15 R-8S</td><td></td></tr><tr><td>Surgical forceps</td><td>Braun Aesculap AG &amp;amp;CoKG, Melsungen, Germany</td><td>BD510R</td><td></td></tr><tr><td>Tape (Heftpflaster Seide) 1.25 cm</td><td>Fink &amp;amp; Walter GmbH, Mechweiler, Germany</td><td>1671801</td><td></td></tr><tr><td>Xylazine&amp;nbsp;</td><td>Bayer Vital GmbH, Leverkusen, Germany</td><td>1320422</td><td></td></tr><tr><td>&amp;alpha;-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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/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.bdigitaluss.remotexs.co/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 ...

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Universidad San Sebastian

Research

JoVE Journal

Bioengineering

12.4K 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

Elastomeric PGS Scaffolds in Arterial Tissue Engineering

Cardiovascular disease is one of the leading cause of mortality in the US and especially, coronary artery disease increases with an aging population and increasing obesity1. Currently, bypass surgery using autologous vessels, allografts, and synthetic grafts are known as a commonly used for arterial substitutes2. However, these grafts have limited applications when an inner diameter of arteries is less than 6 mm due to low availability, thrombotic complications, compliance mismatch, and late intimal hyperplasia3,4. To overcome these limitations, tissue engineering has been successfully applied as a promising alternative to develop small-diameter arterial constructs that are nonthrombogenic, robust, and compliant. Several previous studies have developed small-diameter arterial constructs with tri-lamellar structure, excellent mechanical properties and burst pressure comparable to native arteries5,6. While high tensile strength and burst pressure by increasing collagen production from a rigid material or cell sheet scaffold, these constructs still had low elastin production and compliance, which is a major problem to cause graft failure after implantation. Considering these issues, we hypothesized that an elastometric biomaterial combined with mechanical conditioning would provide elasticity and conduct mechanical signals more efficiently to vascular cells, which increase extracellular matrix production and support cellular orientation.
The objective of this report is to introduce a fabrication technique of porous tubular scaffolds and a dynamic mechanical conditioning for applying them to arterial tissue engineering. We used a biodegradable elastomer, poly (glycerol sebacate) (PGS)7 for fabricating porous tubular scaffolds from the salt fusion method. Adult primary baboon smooth muscle cells (SMCs) were seeded on the lumen of scaffolds, which cultured in our designed pulsatile flow bioreactor for 3 weeks. PGS scaffolds had consistent thickness and randomly distributed macro- and micro-pores. Mechanical conditioning from pulsatile flow bioreactor supported SMC orientation and enhanced ECM production in scaffolds. These results suggest that elastomeric scaffolds and mechanical conditioning of bioreactor culture may be a promising method for arterial tissue engineering.

Elastomeric PGS scaffolds with vascular smooth muscle cells cultured in a pulsatile flow bioreactor may lead to promising small-diameter arterial constructs with native ECM production in a relatively short culture period.

Cardiovascular disease is one of the leading cause of mortality in the US and especially, coronary artery disease increases with an aging population and ...

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 ...

Three-Dimensional Culture of Vascularized Thermogenic Adipose Tissue from Microvascular Fragments

Engineering thermogenic adipose tissue (e.g., beige or brown adipose tissues) has been investigated as a potential therapy for metabolic diseases or for the design of personalized microtissues for health screening and drug testing. Current strategies are often quite complex and fail to accurately fully depict the multicellular and functional properties of thermogenic adipose tissue. Microvascular fragments, small intact microvessels comprised of arteriole, venules, and capillaries isolated from adipose tissue, serve as a single autologous source of cells that enable vascularization and adipose tissue formation. This article describes methods for optimizing culture conditions to enable the generation of three-dimensional, vascularized, and functional thermogenic adipose tissues from microvascular fragments, including protocols for isolating microvascular fragments from adipose tissue and culture conditions. Additionally, best practices are discussed, as are techniques for characterizing the engineered tissues, and sample results from both rodent and human microvascular fragments are provided. This approach has the potential to be utilized for the understanding and development of treatments for obesity and metabolic disease.

Here, we present a detailed protocol outlining the use of microvascular fragments isolated from rodent or human fat tissue as a straightforward approach to engineer functional, vascularized beige adipose tissue.

Engineering thermogenic adipose tissue (e.g., beige or brown adipose tissues) has been investigated as a potential therapy for metabolic diseases or for ...

Human Adipose Tissue Micro-fragmentation for Cell Phenotyping and Secretome Characterization

In the past decade, adipose tissue transplants have been widely used in plastic surgery and orthopaedics to enhance tissue repletion and/or regeneration. Accordingly, techniques for harvesting and processing human adipose tissue have evolved in order to quickly and efficiently obtain large amounts of tissue. Among these, the closed system technology represents an innovative and easy-to-use system to harvest, process, and re-inject refined fat tissue in a short time and in the same intervention (intra-operatively). Adipose tissue is collected by liposuction, washed, emulsified, rinsed and minced mechanically into cell clusters of 0.3 to 0.8 mm. Autologous transplantation of mechanically fragmented adipose tissue has shown remarkable efficacy in different therapeutic indications such as aesthetic medicine and surgery, orthopedic and general surgery. Characterization of micro-fragmented adipose tissue revealed the presence of intact small vessels within the adipocyte clusters; hence, the perivascular niche is left unperturbed. These clusters are enriched in perivascular cells (i.e., mesenchymal stem cell (MSC) ancestors) and in vitro analysis showed an increased release of growth factors and cytokines involved in tissue repair and regeneration, compared to enzymatically derived MSCs. This suggests that the superior therapeutic potential of microfragmented adipose tissue is explained by a higher frequency of presumptive MSCs and enhanced secretory activity. Whether these added pericytes directly contribute to higher growth factor and cytokine production is not known. This clinically approved procedure allows the transplantation of presumptive MSCs without the need for expansion and/or enzymatic treatment, thus bypassing the requirements of GMP guidelines, and reducing the costs for cell-based therapies.

Here, we present human adipose tissue enzyme-free micro-fragmentation using a closed system device. This new method allows the obtainment of sub-millimeter clusters of adipose tissue suitable for in vivo transplantation, in vitro culture, and further cell isolation and characterization.

In the past decade, adipose tissue transplants have been widely used in plastic surgery and orthopaedics to enhance tissue repletion and/or regeneration. ...

Biology

Isolation and Magnetic Separation of Murine CD34+ Stem Cells from the Vascular Wall

Immunomagnetic Isolation of the Vascular Wall-Resident CD34+ Stem Cells from Mice

Resident CD34+ vascular wall-resident stem and progenitor cells (VW-SCs) are increasingly recognized for their crucial role in regulating vascular injury and repair. Establishing a stable and efficient method to culture functional murine CD34+ VW-SCs is essential for further investigating the mechanisms involved in the proliferation, migration, and differentiation of these cells under various physiological and pathological conditions. The described method combines magnetic bead screening and flow cytometry to purify primary cultured resident CD34+ VW-SCs. The purified cells are then functionally identified through immunofluorescence staining and Ca2+ imaging. Briefly, vascular cells from the adventitia of the murine aorta and mesenteric artery are obtained through tissue block attachment, followed by subculturing until reaching a cell count of at least 1 &#215; 107. Subsequently, CD34+ VW-SCs are purified using magnetic bead sorting and flow cytometry. Identification of CD34+ VW-SCs involves cellular immunofluorescence staining, while functional multipotency is determined by exposing cells to a specific culture medium for oriented differentiation. Moreover, functional internal Ca2+ release and external Ca2+ entry is assessed using a commercially available imaging workstation in Fura-2/AM-loaded cells exposed to ATP, caffeine, or thapsigargin (TG). This method offers a stable and efficient technique for isolating, culturing, and identifying vascular wall-resident CD34+ stem cells, providing an opportunity for in vitro studies on the regulatory mechanisms of VW-SCs and the screening of targeted drugs.

Author Spotlight: Enhanced Isolation and Characterization of Vascular Wall Resident Murine CD34+ Stem Cells Using Magnetic Bead Screening-Coupled Flow Cytometry

This study has established a stable and efficient method for the isolation, culture, and functional determination of vascular wall-resident CD34+ stem cells (CD34+ VW-SCs). This easy-to-follow and time-effective isolation method can be utilized by other investigators to study the potential mechanisms involved in cardiovascular diseases.

Resident CD34+ vascular wall-resident stem and progenitor cells (VW-SCs) are increasingly recognized for their crucial role in regulating vascular injury ...

Medicine

Isolation and Identification of Vascular Endothelial Cells from Distinct Adipose Depots for Downstream Applications

Vascular endothelial cells lining the wall of the vascular system play important roles in a variety of physiological processes, including vascular tone regulation, barrier functions, and angiogenesis. Endothelial cell dysfunction is a hallmark predictor and major driver for the progression of severe cardiovascular diseases, yet the underlying mechanisms remain poorly understood. The ability to isolate and perform analyses on endothelial cells from various vascular beds in their native form will give insight into the processes of cardiovascular disease. This protocol presents the procedure for the dissection of mouse subcutaneous and mesenteric adipose tissues, followed by isolation of their respective arterial vasculature. The isolated arteries are then digested using a specific cocktail of digestive enzymes focused on liberating functionally viable endothelial cells. The digested tissue is assessed by flow cytometry analysis using CD31+/CD45&#8722; cells as markers for positive endothelial cell identification. Cells can be sorted for immediate downstream functional assays or used to generate primary cell lines. The technique of isolating and digesting arteries from different vascular beds will provide options for researchers to evaluate freshly isolated vascular cells&#160;from arteries of interest and allow them to perform a wide range of functional tests on specific cell types.

This protocol details a method for the dissection of mouse adipose depots and the isolation and digestion of respective arteries to liberate and then identify the endothelial cell population. Freshly isolated cells used in downstream applications will advance the understanding of vascular cell biology and the mechanisms of vascular dysfunction.

Vascular endothelial cells lining the wall of the vascular system play important roles in a variety of physiological processes, including vascular tone ...

Semi-Automated Isolation of Murine White Adipose Tissue

Semi-Automated Isolation of the Stromal Vascular Fraction from Murine White Adipose Tissue Using a Tissue Dissociator

The in vitro study of white, brown, and beige adipocyte differentiation enables the investigation of cell-autonomous functions of adipocytes and their mechanisms. Immortalized white preadipocyte cell lines are publicly available and widely used. However, the emergence of beige adipocytes in white adipose tissue in response to external cues is difficult to recapitulate to the full extent using publicly available white adipocyte cell lines. Isolation of the stromal vascular fraction (SVF) from murine adipose tissue is commonly executed to obtain primary preadipocytes and perform adipocyte differentiation. However, mincing and collagenase digestion of adipose tissue by hand can result in experimental variation and is prone to contamination. Here, we present a modified semi-automated protocol that utilizes a tissue dissociator for collagenase digestion to achieve easier isolation of the SVF, with the aim of reducing experimental variation, reducing contamination, and increasing reproducibility. The obtained preadipocytes and differentiated adipocytes can be used for functional and mechanistic analyses.

Author Spotlight: Semi-Automated Isolation of the Stromal Vascular Fraction from Murine White Adipose Tissue Using a Tissue Dissociator  

This protocol describes the semi-automated isolation of the stromal vascular fraction (SVF) from murine adipose tissue to obtain preadipocytes and achieve adipocyte differentiation in vitro. Using a tissue dissociator for collagenase digestion reduces experimental variation and increases reproducibility.

The in vitro study of white, brown, and beige adipocyte differentiation enables the investigation of cell-autonomous functions of adipocytes and their ...

Culturing Preadipocytes Isolated from Murine Perivascular Adipose Tissue

Isolation, Culture, and Adipogenic Induction of Stromal Vascular Fraction-derived Preadipocytes from Mouse Periaortic Adipose Tissue

Perivascular adipose tissue (PVAT) is an adipose tissue depot that surrounds blood vessels and exhibits the phenotypes of white, beige, and brown adipocytes. Recent discoveries have shed light on the central role of PVAT in regulating vascular homeostasis and participating in the pathogenesis of cardiovascular diseases. A comprehensive understanding of PVAT properties and regulation is of great importance for the development of future therapies. Primary cultures of periaortic adipocytes are valuable for studying PVAT function and the crosstalk between periaortic adipocytes and vascular cells. This paper presents an economical and feasible protocol for the isolation, culture, and adipogenic induction of stromal vascular fraction-derived preadipocytes from mouse periaortic adipose tissue, which can be useful for modeling adipogenesis or lipogenesis in vitro. The protocol outlines tissue processing and cell differentiation for culturing periaortic adipocytes from young mice. This protocol will provide the technological cornerstone at the bench side for the investigation of PVAT function.

Author Spotlight: The Significance of Isolation, Culture, and Adipogenic Induction of SVF-Derived Preadipocytes from Mouse Perivascular Adipose Tissue 

Here, we describe the isolation, culture, and adipogenic induction of stromal vascular fraction-derived preadipocytes from mouse periaortic adipose tissue, allowing for the study of perivascular adipose tissue function and its relationship with vascular cells.

Perivascular adipose tissue (PVAT) is an adipose tissue depot that surrounds blood vessels and exhibits the phenotypes of white, beige, and brown ...

Isolating Adipose-Derived Stem Cells from Murine Periaortic Adipose Tissue

Source: Qi, Y., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/60691">Isolation, Culture, and Adipogenic Induction of Neural Crest Original Adipose-Derived Stem Cells from Periaortic Adipose Tissue.</a> J. Vis. Exp. (2020)
This video demonstrates a technique for isolating and culturing fluorescently labeled neural crest-derived adipose-derived stem cells (ADSCs) from murine periaortic arch adipose tissue. Following isolation, the NCADSCs are detected, sorted using fluorescence-activated cell sorting, and subsequently cultured on a plate, promoting cell proliferation.

Izolowanie komórek macierzystych pochodzących z tkanki tłuszczowej myszy okołoortowej

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Isolation of the stromal vascular ...