We thank Danielle Berger, Marilyn Woodruff, Simone Correa, and Retha Geiss for assistance with study coordination. SEM was performed by University of Michigan Microscopy &#38; Image Analysis Laboratory Biomedical Research Core Facility. This project was supported by NIH grants R01DK097449 (RWO), R01DK115190 (RWO, CNL), R01DK090262 (CNL), Veterans Affairs Merit Grant I01CX001811 (RWO), Pilot and Feasibility Grant from the Michigan Diabetes Research Center (NIH Grant P30-DK020572) (RWO), Veterans Administration VISN 10 SPARK Pilot Grant (RWO). Scanning electron microscopy performed by the University of Michigan Microscopy &#38; Image Analysis Laboratory Biomedical Research Core Facility. Figure 4 of this manuscript was originally published in Baker et al., J Clin Endo Metab 2017; Mar 1;102 (3), 1032-1043. doi: 10.1210/jc.2016-2915, and has been reproduced by permission of Oxford University Press [https://academic-oup-com.remotexs.ntu.edu.sg/jcem/article/102/3/1032/2836329]. For permission to reuse this material, please visit http://global.oup.com/academic/rights.

<ol>
	<li>Frantz, C., Stewart, K. M., Weaver, V. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+extracellular+matrix+at+a+glance.">The extracellular matrix at a glance.</a> Journal of Cell Science. 123, 4195-4200 (2010).</li><li>Berrier, A. L., Yamada, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell-matrix+adhesion.">Cell-matrix adhesion.</a> Journal of Cell Physiology. 213 (3), 565-573 (2007).</li><li>Trayhurn, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hypoxia+and+adipose+tissue+function+and+dysfunction+in+obesity.">Hypoxia and adipose tissue function and dysfunction in obesity.</a> Physiology Reviews. 93 (1), 1-21 (2014).</li><li>O'Rourke, R. W., Lumeng, C. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Obesity+heats+up+adipose+tissue+lymphocytes.">Obesity heats up adipose tissue lymphocytes.</a> Gastroenterology. 145 (2), 282-285 (2013).</li><li>Engin, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Pathogenesis+of+Obesity-Associated+Adipose+Tissue+Inflammation.">The Pathogenesis of Obesity-Associated Adipose Tissue Inflammation.</a> Advances in Experimental Medicine and Biology. 960. 960, 221-245 (2017).</li><li>Dankel, S. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=COL6A3+expression+in+adipocytes+associates+with+insulin+resistance+and+depends+on+PPAR&#947;+and+adipocyte+size.">COL6A3 expression in adipocytes associates with insulin resistance and depends on PPAR&#947; and adipocyte size.</a> Obesity (Silver Spring). 22 (8), 1807-1813 (2014).</li><li>Divoux, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fibrosis+in+human+adipose+tissue:+composition,+distribution,+and+link+with+lipid+metabolism+and+fat+mass+loss.">Fibrosis in human adipose tissue: composition, distribution, and link with lipid metabolism and fat mass loss.</a> Diabetes. 59, 2817-2825 (2010).</li><li>Lackey, D. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contributions+of+adipose+tissue+architectural+and+tensile+properties+toward+defining+healthy+and+unhealthy+obesity.">Contributions of adipose tissue architectural and tensile properties toward defining healthy and unhealthy obesity.</a> American Journal of Physiology, Endocrinology, and Metabolism. 306 (3), E233-E246 (2014).</li><li>Muir, L. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipose+tissue+fibrosis,+hypertrophy,+and+hyperplasia:+correlations+with+diabetes+in+human+obesity.">Adipose tissue fibrosis, hypertrophy, and hyperplasia: correlations with diabetes in human obesity.</a> Obesity (Silver Spring). 24 (3), 597-605 (2016).</li><li>Spencer, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adipose+tissue+macrophages+in+insulin-resistant+subjects+are+associated+with+collagen+VI+and+fibrosis+and+demonstrate+alternative+activation.">Adipose tissue macrophages in insulin-resistant subjects are associated with collagen VI and fibrosis and demonstrate alternative activation.</a> American Journal of Physiology, Endocrinology, and Metabolism. 299 (6), E1016-E1027 (2010).</li><li>Baker, N. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diabetes-specific+regulation+of+adipocyte+metabolism+by+the+adipose+tissue+extracellular+matrix.">Diabetes-specific regulation of adipocyte metabolism by the adipose tissue extracellular matrix.</a> Journal of Clinical Endocrinology and Metabolism. 102 (3), 1-12 (2017).</li><li>Pellegrinelli, V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+adipocyte+function+is+impacted+by+mechanical+cues.">Human adipocyte function is impacted by mechanical cues.</a> Journal of Patholology. 233 (2), 183-195 (2014).</li><li>Flynn, L. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+decellularized+adipose+tissue+to+provide+an+inductive+microenvironment+for+the+adipogenic+differentiation+of+human+adipose-derived+stem+cells.">The use of decellularized adipose tissue to provide an inductive microenvironment for the adipogenic differentiation of human adipose-derived stem cells.</a> Biomaterials. 31 (17), 4715-4724 (2010).</li><li>Perea-Gil, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+comparative+study+of+two+decellularization+protocols+in+search+of+an+optimal+myocardial+scaffold+for+recellularization.">In vitro comparative study of two decellularization protocols in search of an optimal myocardial scaffold for recellularization.</a> American Journal Translational Research. 7 (3), 558-573 (2015).</li><li>Porzionato, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Decellularized+omentum+as+novel+biologic+scaffold+for+reconstructive+surgery+and+regenerative+medicine.">Decellularized omentum as novel biologic scaffold for reconstructive surgery and regenerative medicine.</a> European Journal of Histochemistry. 57 (1), e4 (2013).</li><li>Tebyanian, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Comparative+Study+of+Rat+Lung+Decellularization+by+Chemical+Detergents+for+Lung+Tissue+Engineering.">A Comparative Study of Rat Lung Decellularization by Chemical Detergents for Lung Tissue Engineering.</a> Open Access Macedonian Journal of Medical Sciences. 5 (7), 859-865 (2017).</li><li>Crapo, P. M., Gilbert, T. W., Badylak, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+overview+of+tissue+and+whole+organ+decellularization+processes.">An overview of tissue and whole organ decellularization processes.</a> Biomaterials. 32 (12), 3233-3243 (2011).</li><li>Wang, L., Johnson, J. A., Zhang, Q., Beahm, E. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Combining+decellularized+human+adipose+tissue+extracellular+matrix+and+adipose-derived+stem+cells+for+adipose+tissue+engineering.">Combining decellularized human adipose tissue extracellular matrix and adipose-derived stem cells for adipose tissue engineering.</a> Acta Biomaterials. 9 (11), 8921-8931 (2013).</li><li>Booth, A. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acellular+normal+and+fibrotic+human+lung+matrices+as+a+culture+system+for+in+vitro+investigation.">Acellular normal and fibrotic human lung matrices as a culture system for in vitro investigation.</a> American Journal of Respiratory and Critical Care Medicine. 186 (9), 866-876 (2012).</li><li>Parker, M. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fibrotic+extracellular+matrix+activates+a+profibrotic+positive+feedback+loop.">Fibrotic extracellular matrix activates a profibrotic positive feedback loop.</a> Journal of Clinical Investigation. 124 (4), 1622-1635 (2014).</li><li>Baker, N. A., Muir, L. A., Lumeng, C. N., O'Rourke, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differentiation+and+Metabolic+Interrogation+of+Human+Adipocytes.">Differentiation and Metabolic Interrogation of Human Adipocytes.</a> Methods in Molecular Biology. 1566, 61-76 (2017).</li><li>O'Rourke, R. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hexosamine+biosynthesis+is+a+possible+mechanism+underlying+hypoxia's+effects+on+lipid+metabolism+in+human+adipocytes.">Hexosamine biosynthesis is a possible mechanism underlying hypoxia's effects on lipid metabolism in human adipocytes.</a> PLoS One. 8 (8), e71165 (2013).</li><li>Tchkonia, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fat+depot-specific+characteristics+are+retained+in+strains+derived+from+single+human+preadipocytes.">Fat depot-specific characteristics are retained in strains derived from single human preadipocytes.</a> Diabetes. 55 (9), 2571-2578 (2006).</li><li>Tchoukalova, Y. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sex-+and+depot-dependent+differences+in+adipogenesis+in+normal-weight+humans.">Sex- and depot-dependent differences in adipogenesis in normal-weight humans.</a> Obesity (Silver Spring). 18 (10), 1875-1880 (2010).</li></ol>

The authors declare no conflicting interests.

The ECM-adipocyte culture model provides a valuable tool for dissecting the individual roles of ECM and cells in dictating ultimate tissue phenotype. The ECM isolation protocol is quite reproducible, but variability in the decellularization process may be observed. The Day 3 delipidation step is a critical point in the protocol. At the completion of the overnight extraction, delipidation of the matrix should be evidenced by the Polar Solvent Solution turning yellow, while the matrix should transition from a yellow-orange color characteristic of intact adipose tissue to translucent/white (Figure 1). If these color changes do not occur, then delipidation is likely not complete. Inter-patient variability in delipidation efficiency is common, but to date we have not observed any correlation of efficiency of delipidation with subject DM status, age, BMI, or other clinical variables. Large samples (greater than 20 g) will not undergo efficient delipidation; processing samples &#60;20 g may prevent this problem. If a sample does not appear completely delipidated after Day 3, divide the sample in half with sterile scissors, then transfer samples to fresh 15-20 mL of Polar Solvent Extraction Solution and place on an orbital shaker for 3-5 h. Some samples may not fully decellularize after repeating the polar solvent extraction step. If a majority of the sample is decellularized, it is possible to cut out the sections containing lipid before using the sample in experiments.
Contamination may occur if sterility is not diligently maintained throughout the process. All work should be performed in a laminar flow hood using standard cell culture precautions. We typically store ECM for up to 3 months at 4 &#176;C, and preadipocytes for up to 6 months in liquid nitrogen. Retention of biofunctionality and disease- and depot-specific properties beyond this time has not been tested.
Confocal microscopy permits visualization of mature adipocytes within the ECM, which may be enhanced with Oil Red-O staining. Scanning electron microscopy (SEM) also provides a non-quantitative method for visualizing adipocytes within ECM. Of note, SEM and Oil Red-O staining suggest unilocular adipocytes in the ECM-adipocyte constructs, a morphology that is similar to in vivo adipocytes and in contrast to the typical multilocular adipocytes observed when preadipocytes undergo adipogenic differentiation on 2D plastic (Figure 3, Figure 4A). This observation suggests that the ECM provides a more physiologic environment for adipogenic differentiation than standard tissue culture on plastic. Fixation and sectioning of ECM-adipocyte constructs may be difficult, as OCT-embedded constructs are fragile and do not section easily. Sectioning embedded tissues immediately after removal from storage from the -80 &#176;C freezer and performing sectioning in a cold-room (4 &#176;C) on a pre-chilled cryostat makes sectioning feasible.
ECM-adipocyte culture provides a physiologic environment that approximates the in vivo adipose tissue environment and provides a sustainable environment for preadipocyte stem cell growth and differentiation, as evidenced by upregulation of adipogenic genes in 3D-ECM-adipocyte culture. ECM-adipocyte cultures also engage in adipocyte metabolic functions, including glucose uptake and lipolysis8. We report glucose uptake as cpm normalized to either total protein extracted from ECM-adipocyte cultures measured with a Bradford assay, or to DNA content measured with a DNA assay kit, and have observed similar results with both methods. Similarly, we report lipolysis as mg/mL of glycerol release normalized to either protein or DNA, also with similar results obtained with either normalization method. Others suggest normalizing adipocyte metabolic data to lipid content, but to date our attempts to reliably extract lipid from ECM constructs has been unsuccessful. Reporting glucose uptake rate as moles of glucose per unit of time and lipolysis as moles of glycerol/free fatty acid per unit of time may permit more accurate comparisons between separate experiments. Isolation of RNA from ECM for qrtPCR analysis is characterized by lower yields compared with cells in 2D culture, but preparation of larger ECM fragments seeded with more cells overcomes this problem, as described in step 8.3.1. We have encountered difficulty in isolating adequate quantities of undegraded protein with phosphomoieties intact for Western blot analysis, which represents a limitation of the model.
Similar methodology for isolating ECM from adipose tissue and seeding with preadipocytes has been previously published, with evidence suggesting that ECM may promote adipogenic differentiation absent classic adipogenic mediators including cAMP agonists, insulin, and PPAR-&#947; agonists12,14. Our data is the first to demonstrate disease-specific regulation of cellular metabolism by ECM in adipose tissue, using methods in which adipocytes are stimulated with classic adipogenic differentiation factors11; similar studies in the absence of adipogenic stimuli are a target of future research. Others have demonstrated similar disease-specific ECM-cell crosstalk using ECM and cells isolated from lung tissue19,20. Alternative strategies have also been employed, including creation of matrices by mixing homogenized adipose tissue ECM preparations in peptide hydrogels12.
While inter-patient variability in human adipocyte cellular metabolism is observed in both 2D-plastic and 3D-ECM culture, when analyzed in aggregate, these cells manifest disease-, depot- and sex-specific differences in cellular metabolism, as described by our group and others11,21,22,23,24, validating the utility and generalizability of these culture models for the study of adipocyte cellular metabolism. We have demonstrated that adipocytes differentiated in disease-matched ECM (i.e., NDM adipocytes in NDM ECM, DM adipocytes in DM ECM) recapitulate metabolic phenotypes of isolated NDM and DM adipocytes in standard 2D culture, supporting ECM-adipocyte culture as a model for studying disease-specific alterations in adipocyte cellular metabolism in a more physiologic environment. We have not observed differences in the magnitude or direction of metabolic functions, including glucose uptake or lipolysis, when ECM and adipocytes from the same or different donors are studied. In either case, ECM-adipocyte constructs manifest DM-specific metabolic phenotype (e.g., decreased GU in DM/DM relative to NDM/NDM constructs), with no clear difference when comparing constructs comprised of ECM and adipocytes from the same or different donors.
ECM-adipocyte culture allows study of combinations of ECM and preadipocytes from distinct patient populations and tissue depots, permitting analysis of disease- and depot-specific contributions of ECM and adipocytes to ultimate adipose tissue metabolic phenotype. Demonstrating this strength of the ECM-adipocyte culture model, we have shown that ECM from NDM tissue has the capacity to rescue metabolic defects in DM adipocytes (Figure 4C)11. Preliminary data in murine visceral and subcutaneous adipose tissues suggest that murine ECM regulates murine adipocyte metabolism similarly in a depot-specific manner (unpublished data, O&#39;Rourke RW, 2019), suggesting that this model system may be used to study ECM-adipocyte crosstalk in both murine and human systems. Furthermore, this model permits study of isolated manipulation of ECM as a means to regulate adipose tissue phenotype. Preliminary study of ECM treated in conditions that induce glycation targeted specifically to the ECM, for example, suggest that these modifications regulate the cellular metabolic phenotype of cells subsequently seeded into treated ECM (unpublished data, O&#39;Rourke RW, 2019), providing a model for study of the effects of targeted manipulation of the ECM on adipocyte phenotype.

The extracellular matrix (ECM) not only provides a mechanical scaffold for tissues, but also engages in complex crosstalk with cells that reside within it, regulating diverse processes necessary for tissue homeostasis, including cell proliferation, differentiation, signaling, and metabolism1. While healthy ECM plays an essential role the maintenance of normal tissue function, dysfunctional ECM has been implicated in multiple diseases2.
Adipose tissue plays an important role in the pathogenesis of metabolic disease. Obesity is associated with excessive adipocyte hypertrophy and cellular hypoxia, defects in adipocyte cellular metabolism, and adipose tissue endoplasmic reticulum and oxidative stress and inflammation. While poorly understood, these complex processes conspire to impair adipose tissue nutrient buffering capacity, leading to nutrient overflow from adipose tissue, toxicity in multiple tissues, and systemic metabolic disease3,4,5.The sequence of events and specific mechanisms that underlie adipose tissue failure are poorly understood, but alterations in adipose tissue ECM have been implicated. The ECM composition is altered within adipose tissue in human and murine obesity, with increased deposition of ECM protein along with qualitative biochemical and structural differences in the adipose tissue ECM associated with human metabolic disease, including type 2 diabetes and hyperlipidemia6,7,8,9,10,11.
Despite these observations, the role of adipose tissue ECM in mediating adipose tissue dysfunction is not well-defined. This is in part due to a lack of tractable experimental models that permit dissection of the specific roles of ECM and adipocytes in regulating ultimate adipose tissue function. ECM-adipocyte culture better simulates the in vivo environment of native adipose tissue in at least two respects. Firstly, ECM culture provides a molecular environment similar to native adipose tissue, including native collagens, elastins, and other matrix proteins absent in standard 2D culture. Secondly, culture on 2D plastic has been shown to alter adipocyte metabolism via mechanical effects due to decreased elasticity of plastic substrate12, which ECM-culture eliminates.
Methods to engineer biological scaffolds by isolation of ECM from decellularized adipose and other tissues have been studied in the context of regenerative and reconstructive medicine and tissue engineering13,14,15,16,17,18. We have previously published methodology in which we adapted these methods to develop an in vitro 3D model of human ECM-adipocyte culture, using ECM and adipocyte stem cells (preadipocytes) derived from human visceral adipose tissues11. In the present article, we describe these methods in detail. The decellularization procedure for human adipose tissue is a four-day process that involves mechanical and enzymatic treatments to remove cells and lipid, leaving a biological scaffold that maintains characteristics of the tissue from which it is derived. Decellularized ECM supports adipogenic differentiation of human preadipocytes, and when reconstituted with adipocytes, maintains microarchitecture and biochemical and disease-specific characteristics of intact adipose tissue and engages in metabolic functions characteristic of native adipose tissue. This matrix can be studied alone or reseeded with cells, permitting study of interactions and crosstalk between the cellular and extra-cellular components of adipose tissue.

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			<td>0.25% trypsin-EDTA</td>
			<td>Gibco, ThermoFisher Scientific Inc., Waltham, MA, USA</td>
			<td>Cat#25200056</td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 mL cryovial tube</td>
			<td>Fisher Scientific, ThermoFisher Scientific Inc., Waltham, MA USA</td>
			<td>Cat#02-682-557</td>
			<td></td>
		</tr>
		<tr>
			<td>10% Neutral Buffered Formalin</td>
			<td>VWR International LLC., Radnor, PA, USA</td>
			<td>Cat#89370-094</td>
			<td></td>
		</tr>
		<tr>
			<td>100 &#181;m nylon mesh filter</td>
			<td>Corning Inc., Corning, NY, USA</td>
			<td>Cat#352360</td>
			<td></td>
		</tr>
		<tr>
			<td>2-Deoxy-D-glucose</td>
			<td>Sigma-Aldrich, Inc., St Louis, MO, USA</td>
			<td>Cat#D8375</td>
			<td></td>
		</tr>
		<tr>
			<td>2 nM 3,3&#8217;-5,Triiodo,L-thyronine sodium salt (T3)</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#T6397</td>
			<td></td>
		</tr>
		<tr>
			<td>24-well tissue culture plates</td>
			<td>VWR International LLC., Radnor, PA, USA</td>
			<td>Cat#10861-700</td>
			<td></td>
		</tr>
		<tr>
			<td>3-Isobutyl-1-methylxanthine (IBMX)</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#I5879</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well tissue culture plates</td>
			<td>VWR International LLC., Radnor, PA, USA</td>
			<td>Cat#10861-666</td>
			<td></td>
		</tr>
		<tr>
			<td>Antibiotic-Antimycotic Solution (ABAM)</td>
			<td>Gibco, ThermoFisher Scientific Inc., Waltham, MA, USA</td>
			<td>Cat#15240062</td>
			<td></td>
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		<tr>
			<td>Biotin</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#B4639</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Sigma-Aldrich, Inc., St Louis, MO, USA</td>
			<td>Cat#A8806</td>
			<td></td>
		</tr>
		<tr>
			<td>Buffer RLT</td>
			<td>Qiagen, Hilden, Germany</td>
			<td>Cat#79216</td>
			<td></td>
		</tr>
		<tr>
			<td>Ciglitizone</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#C3974</td>
			<td></td>
		</tr>
		<tr>
			<td>Deoxy-D-glucose, 2-[1,2-3H (N)]-</td>
			<td>PerkinElmer Inc., Waltham, MA, USA</td>
			<td>Cat#NET328A250UC</td>
			<td></td>
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			<td>Deoxyribonuclease I from bovine pancreas, type II-S</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#D4513</td>
			<td></td>
		</tr>
		<tr>
			<td>Dexamethasone</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#D4902</td>
			<td></td>
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		<tr>
			<td>Dimethyl Sulfoxide</td>
			<td>Fisher Scientific, ThermoFisher Scientific Inc., Waltham, MA USA</td>
			<td>Cat#BP231</td>
			<td>Flammable, caustic</td>
		</tr>
		<tr>
			<td>Disodium EDTA</td>
			<td>Fisher Scientific, ThermoFisher Scientific Inc., Waltham, MA USA</td>
			<td>Cat#BP118</td>
			<td></td>
		</tr>
		<tr>
			<td>D-pantothenic acid hemicalcium salt</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#21210</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s Modified Eagle Medium: Nutrient Mixture F-12 (DMEM/F12</td>
			<td>Gibco, ThermoFisher Scientific Inc., Waltham, MA USA</td>
			<td>Cat#11320033</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Decon Labs, Inc., King of Prussia, PA, USA</td>
			<td>Cat#DSP-MD.43</td>
			<td>Flammable</td>
		</tr>
		<tr>
			<td>EVE Cell Counting Slides, NanoEnTek</td>
			<td>VWR International LLC., Radnor, PA, USA</td>
			<td>Cat#10027-446</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>Gibco, ThermoFisher Scientific Inc., Waltham, MA, USA</td>
			<td>Cat#10437028</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde</td>
			<td>Sigma-Aldrich, Inc., St Louis, MO, USA</td>
			<td>Cat#G5882</td>
			<td>Caustic</td>
		</tr>
		<tr>
			<td>Hexamethyldisalizane</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#440191</td>
			<td>Flammable, caustic</td>
		</tr>
		<tr>
			<td>Human insulin solution</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#I9278</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropanol</td>
			<td>Fisher Scientific, ThermoFisher Scientific Inc., Waltham, MA USA</td>
			<td>Cat#A415</td>
			<td>Flammable</td>
		</tr>
		<tr>
			<td>Isoproterenol</td>
			<td>Sigma-Aldrich, Inc., St Louis, MO, USA</td>
			<td>Cat#I5627</td>
			<td>Flammable</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#S25484</td>
			<td></td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#P5655</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipase from porcine pancreas, type VI-S</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#L0382</td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4*7H2O</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#230391</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#S5136</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#S3014</td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Fisher Scientific, ThermoFisher Scientific Inc., Waltham, MA USA</td>
			<td>Cat#S233</td>
			<td></td>
		</tr>
		<tr>
			<td>NH4Cl</td>
			<td>Fisher Scientific, ThermoFisher Scientific Inc., Waltham, MA USA</td>
			<td>Cat#A661</td>
			<td></td>
		</tr>
		<tr>
			<td>Optimal cutting temperature (OCT) compound</td>
			<td>Agar Scientific, Ltd., Stansted, Essex, UK</td>
			<td>Cat# AGR1180</td>
			<td></td>
		</tr>
		<tr>
			<td>Oil Red-O Solution (ORO)</td>
			<td>Sigma-Aldrich, Inc., St Louis, MO, USA</td>
			<td>Cat#O1391</td>
			<td></td>
		</tr>
		<tr>
			<td>Oil Red-O Stain Kit</td>
			<td>American Master Tech Scientific Inc., Lodi, CA, USA</td>
			<td>Cat#KTORO-G</td>
			<td></td>
		</tr>
		<tr>
			<td>Osmium tetroxide</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#201030</td>
			<td>Caustic</td>
		</tr>
		<tr>
			<td>Phenylmethylsulfonyl fluoride (PMSF)</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#93482</td>
			<td>Caustic</td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline Solution (PBS)</td>
			<td>Fisher Scientific, ThermoFisher Scientific Inc., Waltham, MA USA</td>
			<td>Cat#SH3025601</td>
			<td></td>
		</tr>
		<tr>
			<td>Ribonuclease A from bovine pancreas, type III-A</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#R5125</td>
			<td></td>
		</tr>
		<tr>
			<td>RNAEasy Fibrous Tissue MiniKit</td>
			<td>Qiagen, Hilden, Germany</td>
			<td>Cat#74704</td>
			<td></td>
		</tr>
		<tr>
			<td>Scintillation Fluid</td>
			<td>Fisher Scientific, ThermoFisher Scientific Inc., Waltham, MA USA</td>
			<td>Cat#SX18</td>
			<td></td>
		</tr>
		<tr>
			<td>Scintillation Counter</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors, forceps, sterile</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sorensen&#39;s phosphate buffer</td>
			<td>Thomas Scientific, Inc., Swedesboro, NJ</td>
			<td>CAS #: 10049-21-5</td>
			<td></td>
		</tr>
		<tr>
			<td>T-150 culture flask</td>
			<td>VWR International LLC., Radnor, PA, USA</td>
			<td>Cat#10062-864</td>
			<td></td>
		</tr>
		<tr>
			<td>TaqMan Gene Expression Master Mix</td>
			<td>ThermoFisher Scientific Inc., Waltham, MA USA</td>
			<td>Cat#4369016</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature-controlled orbital shaker</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Homogenizer, BeadBug Microtube Homogenizer</td>
			<td>Benchmark Scientific</td>
			<td>Cat#D1030</td>
			<td></td>
		</tr>
		<tr>
			<td>Transferrin</td>
			<td>Sigma-Aldrich, Inc. St Louis, MO, USA</td>
			<td>Cat#T3309</td>
			<td></td>
		</tr>
		<tr>
			<td>Triglyceride Determination Kit</td>
			<td>Sigma-Aldrich, Inc., St Louis, MO, USA</td>
			<td>Cat#TR0100</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan blue stain, 0.4%</td>
			<td>VWR International LLC., Radnor, PA, USA</td>
			<td>Cat#10027-446</td>
			<td></td>
		</tr>
		<tr>
			<td>Type II collagenase</td>
			<td>Gibco, ThermoFisher Scientific Inc., Waltham, MA, USA</td>
			<td>Cat#17101015</td>
			<td></td>
		</tr>
		<tr>
			<td>Whatman Reeve Angel filter paper, Grade 201, 150mm</td>
			<td>Sigma-Aldrich, Inc., St Louis, MO, USA</td>
			<td>Cat#WHA5201150</td>
			<td></td>
		</tr>
	</tbody>
</table>

a human 3d extracellular matrix-adipocyte culture model for studying matrix-cell metabolic crosstalk

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of cellular function. The ECM plays a particularly important role in adipose tissue function in obesity, and alterations in adipose tissue ECM deposition and composition are associated with metabolic disease in mice and humans. Tractable in vitro models that permit dissection of the roles of the ECM and cells in contributing to global tissue phenotype are sparse. We describe a novel 3D in vitro model of human ECM-adipocyte culture that permits study of the specific roles of the ECM and adipocytes in regulating adipose tissue metabolic phenotype. Human adipose tissue is decellularized to isolate ECM, which is subsequently repopulated with preadipocytes that are then differentiated within the ECM into mature adipocytes. This method creates ECM-adipocyte constructs that are metabolically active and retain characteristics of the tissues and patients from which they are derived. We have used this system to demonstrate disease-specific ECM-adipocyte crosstalk in human adipose tissue. This culture model provides a tool for dissecting the roles of the ECM and adipocytes in contributing to global adipose tissue metabolic phenotype and permits study of the role of the ECM in regulating adipose tissue homeostasis.

Preparation of adipose tissue ECM, seeding with preadipocytes, and in vitro differentiation into mature adipocytes result in clear sequential morphologic changes in tissue that permits visual assessment of progress throughout the protocol (Figure 1). Preadipocytes used to seed the ECM are isolated using collagenase digestion from separate VAT samples (Figure 2). Scanning electron microscopy of ECM-adipocyte constructs at each stage of processing reveals decellularization of ECM and subsequent recapitulation of lipid-containing adipocytes upon reseeding and differentiation (Figure 3). Collagen 1-specific immunohistochemistry of decellularized ECM reveals maintenance of collagen microarchitecture, while Oil Red-O staining of live and fixed/sectioned 3D ECM-adipocyte constructs reveals lipid-containing adipocytes (Figure 4A). qrtPCR from adipocytes differentiated in ECM demonstrates marked upregulation of adipogenic genes relative to undifferentiated preadipocytes cultured in ECM (Figure 4B). Metabolic phenotyping of ECM-adipocyte constructs studying all combinations of ECM and adipocytes from diabetic (DM) and non-diabetic (NDM) subjects reveals impaired glucose uptake and lipolytic function in ECM-adipocyte constructs from DM relative to NDM tissues, recapitulating DM-specific metabolic defects we have previously reported in human adipocytes in standard 2D culture11. Importantly, NDM ECM rescues insulin-stimulated glucose uptake and lipolytic capacity in DM adipocytes&#160;(Figure 4C)11. Together, these data demonstrate disease-specific regulation of adipocyte cellular metabolism by the ECM. Further details are reported in Baker et al.11.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60486/60486fig1.jpg" /> 
Figure 1: Workflow for ECM-adipocyte preparation. Day 1: Whole visceral adipose tissue samples undergo three freeze-thaw cycles followed by overnight incubation with enzymatic digestion solution #1. Day 2: Following digestion with enzymatic solution #1, samples are digested overnight with enzymatic digestion solution #2. At this point, samples will be partially delipidated. Day 3: Following enzymatic digestion #2, samples undergo overnight incubation with polar solvent extraction solution, which will complete the delipidation. After Day 3, samples should be fully delipidated and white/translucent in color. Samples are washed thoroughly with rinsing buffer solution then 70% ethanol and stored in storage solution in 40 &#176;C until ready for reseeding with preadipocytes. Day 4: Preadipocytes are seeded into decellularized VAT followed by adipogenic differentiation for 14 days. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60486/60486fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60486/60486fig2.jpg" /> 
Figure 2: Workflow for preadipocyte isolation. VAT is minced, digested with collagenase, filtered, centrifuged, and the resultant stromovascular cell pellet plated and cultured to expand preadipocytes. See Baker et al. 201721 for further details regarding human preadipocyte isolation and 2D adipocyte culture. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60486/60486fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60486/60486fig3.jpg" /> 
Figure 3: Scanning electron microscopy images. Intact adipose tissue, decellularized adipose tissue, and decellularized adipose tissue repopulated with preadipocytes and differentiated in adipogenic media for 14 days. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60486/60486fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60486/60486fig4.jpg" /> 
Figure 4: Characterization of adipocytes in ECM. (A) Decellularized adipose tissue ECM maintains microarchitecture and supports adipocyte differentiation: Top: Collagen 1 immunohistochemistry of whole human VAT before and after decellularization demonstrating maintenance of microarchitecture; middle: 3D confocal photomicrographs of live human adipocytes within ECM; intact decellularized human VAT stained with Oil Red-O before reseeding with preadipocytes, 4 days after seeding, and 14 days after seeding with preadipocytes followed by adipogenic differentiation; blue: DAPI staining of cell nuclei; red: Oil Red-O staining of intracellular lipid; bottom: Formalin-fixed, paraffin-embedded, 5 &#181;m sectioned, Oil Red-O-stained human VAT prior to decellularization, immediately after decellularization, and after decellularization, preadipocyte-seeding, and 14 days of adipogenic differentiation, demonstrating cytoplasmic lipid accumulation in adipocytes within ECM. (B) Adipocytes in ECM upregulate adipogenic gene expression: qrtPCR analysis comparing adipogenic gene transcript levels in RNA from human VAT adipocytes differentiated in VAT ECM for 14 days relative to undifferentiated preadipocytes in VAT ECM cultured for 72 h in nonadipogenic media. Data are mean &#177; SEM. ACLY: ATP citrate lyase, ATGL: adipose triglyceride lipase, FASN: fatty acid synthase, PPARg: peroxisome proliferative activated receptor gamma; ordinate: fold difference in transcript level in mature adipocytes relative to undifferentiated preadipocyte referent=1; all fold differences significant (p&#60;0.001, paired t-test); ECM from 10 subjects, preadipocytes/adipocytes from n=11 subjects. (C) ECM regulates adipocyte metabolism in a DM-specific manner: 3D-ECM from DM or NDM subjects seeded with preadipocytes from DM or NDM subjects, differentiated into adipocytes, and studied with metabolic phenotyping. Data bars labeled with patient source (NDM, DM) of ECM and adipocytes (ECM/AD); e.g., NDM/NDM denotes both ECM and preadipocytes derived from NDM patients, while NDM/DM denotes ECM from NDM patients combined with preadipocytes from DM patients. Data are mean &#177; SEM. Ordinates: glucose uptake (cpm), normalized to ECM/cell lysate protein concentration(mg/mL); lipolysis: culture supernatant glycerol concentration(mg/mL) normalized to ECM/cell lysate DNA concentration(ng/mL); *p&#60;0.050, **p&#60;0.100 comparing indicated data-point to corresponding data-point (basal or insulin-stimulated for glucose uptake, basal or isoproterenol-stimulated for lipolysis) in DM/DM arm, using mixed model analysis adjusting for repeated measures; ECM from n=9 NDM, 8 DM subjects, preadipocytes from 10 NDM, 9 DM subjects for glucose uptake; ECM from 6 NDM, 6 DM subjects, preadipocytes from 6 NDM, 6 DM subjects for lipolysis. This figure has been adapted from O&#39;Rourke and Lumeng11. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60486/60486fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Human 3D Extracellular Matrix-Adipocyte Culture Model for Studying Matrix-Cell Metabolic Crosstalk at JoVE.com

A Human 3D Extracellular Matrix-Adipocyte Culture Model for Studying Matrix-Cell Metabolic Crosstalk

We describe a 3D human extracellular matrix-adipocyte in vitro culture system that permits dissection of the roles of the matrix and adipocytes in contributing to adipose tissue metabolic phenotype.

The extracellular matrix (ECM) plays a central role in regulating tissue homeostasis, engaging in crosstalk with cells and regulating multiple aspects of ...

a-human-3d-extracellular-matrix-adipocyte-culture-model-for-studying

Nanyang Technological University

Research

JoVE Journal

Bioengineering

9.2K Views.  University of Michigan Medical School. We describe a 3D human extracellular matrix-adipocyte in vitro culture system that permits dissection of the roles of the matrix and adipocytes in contributing to adipose tissue metabolic phenotype.

Video: A Human 3D Extracellular Matrix-Adipocyte Culture Model for Studying Matrix-Cell Metabolic Crosstalk

A Simple Hanging Drop Cell Culture Protocol for Generation of 3D Spheroids

Studies of cell-cell cohesion and cell-substratum adhesion have historically been performed on monolayer cultures adherent to rigid substrates. Cells within a tissue, however, are typically encased within a closely packed tissue mass in which cells establish intimate connections with many near-neighbors and with extracellular matrix components. Accordingly, the chemical milieu and physical forces experienced by cells within a 3D tissue are fundamentally different than those experienced by cells grown in monolayer culture. This has been shown to markedly impact cellular morphology and signaling. Several methods have been devised to generate 3D cell cultures including encapsulation of cells in collagen gels1or in biomaterial scaffolds2. Such methods, while useful, do not recapitulate the intimate direct cell-cell adhesion architecture found in normal tissues. Rather, they more closely approximate culture systems in which single cells are loosely dispersed within a 3D meshwork of ECM products. Here, we describe a simple method in which cells are placed in hanging drop culture and incubated under physiological conditions until they form true 3D spheroids in which cells are in direct contact with each other and with extracellular matrix components. The method requires no specialized equipment and can be adapted to include addition of any biological agent in very small quantities that may be of interest in elucidating effects on cell-cell or cell-ECM interaction. The method can also be used to co-culture two (or more) different cell populations so as to elucidate the role of cell-cell or cell-ECM interactions in specifying spatial relationships between cells. Cell-cell cohesion and cell-ECM adhesion are the cornerstones of studies of embryonic development, tumor-stromal cell interaction in malignant invasion, wound healing, and for applications to tissue engineering. This simple method will provide a means of generating tissue-like cellular aggregates for measurement of biomechanical properties or for molecular and biochemical analysis in a physiologically relevant model.

We describe a simple, rapid method of generating 3D tissue-like spheroids and their potential application to quantify differences in cell-cell interactions.

Studies of cell-cell cohesion and cell-substratum adhesion have historically been performed on monolayer cultures adherent to rigid substrates. Cells ...

Rotating Cell Culture Systems for Human Cell Culture: Human Trophoblast Cells as a Model

The field of human trophoblast research aids in understanding the complex environment established during placentation. Due to the nature of these studies, human in vivo experimentation is impossible. A combination of primary cultures, explant cultures and trophoblast cell lines1 support our understanding of invasion of the uterine wall2 and remodeling of uterine spiral arteries3,4 by extravillous trophoblast cells (EVTs), which is required for successful establishment of pregnancy. Despite the wealth of knowledge gleaned from such models, it is accepted that in vitro cell culture models using EVT-like cell lines display altered cellular properties when compared to their in vivo counterparts5,6. Cells cultured in the rotating cell culture system (RCCS) display morphological, phenotypic, and functional properties of EVT-like cell lines that more closely mimic differentiating in utero EVTs, with increased expression of genes mediating invasion (e.g. matrix metalloproteinases (MMPs)) and trophoblast differentiation7,8,9. The Saint Georges Hospital Placental cell Line-4 (SGHPL-4) (kindly donated by Dr. Guy Whitley and Dr. Judith Cartwright) is an EVT-like cell line that was used for testing in the RCCS.
The design of the RCCS culture vessel is based on the principle that organs and tissues function in a three-dimensional (3-D) environment. Due to the dynamic culture conditions in the vessel, including conditions of physiologically relevant shear, cells grown in three dimensions form aggregates based on natural cellular affinities and differentiate into organotypic tissue-like assemblies10,11,12 . The maintenance of a fluid orbit provides a low-shear, low-turbulence environment similar to conditions found in vivo. Sedimentation of the cultured cells is countered by adjusting the rotation speed of the RCCS to ensure a constant free-fall of cells. Gas exchange occurs through a permeable hydrophobic membrane located on the back of the bioreactor. Like their parental tissue in vivo, RCCS-grown cells are able to respond to chemical and molecular gradients in three dimensions (i.e. at their apical, basal, and lateral surfaces) because they are cultured on the surface of porous microcarrier beads. When grown as two-dimensional monolayers on impermeable surfaces like plastic, cells are deprived of this important communication at their basal surface. Consequently, the spatial constraints imposed by the environment profoundly affect how cells sense and decode signals from the surrounding microenvironment, thus implying an important role for the 3-D milieu13.
We have used the RCCS to engineer biologically meaningful 3-D models of various human epithelial tissues7,14,15,16. Indeed, many previous reports have demonstrated that cells cultured in the RCCS can assume physiologically relevant phenotypes that have not been possible with other models10,17-21. In summary, culture in the RCCS represents an easy, reproducible, high-throughput platform that provides large numbers of differentiated cells that are amenable to a variety of experimental manipulations. 
In the following protocol, using EVTs as an example, we clearly describe the steps required to three-dimensionally culture adherent cells in the RCCS.

Traditional, two dimensional cell culture techniques often result in altered characteristics with respect to differentiation markers, cytokines and growth factors. Three-dimensional cell culture in the rotating cell culture system (RCCS) reestablishes expression of many of these factors as shown here with an extravillous trophoblast cell line.

The field of human trophoblast research aids in  understanding the complex environment established during placentation. Due to the  nature of these ...

Microfluidic Mixers for Studying Protein Folding

The process by which a protein folds into its native conformation is highly relevant to biology and human health yet still poorly understood. One reason for this is that folding takes place over a wide range of timescales, from nanoseconds to seconds or longer, depending on the protein1. Conventional stopped-flow mixers have allowed measurement of folding kinetics starting at about 1 ms. We have recently developed a microfluidic mixer that dilutes denaturant ~100-fold in ~8 &mu;s2. Unlike a stopped-flow mixer, this mixer operates in the laminar flow regime in which turbulence does not occur. The absence of turbulence allows precise numeric simulation of all flows within the mixer with excellent agreement to experiment3-4.
Laminar flow is achieved for Reynolds numbers Re &le;100. For aqueous solutions, this requires micron scale geometries. We use a hard substrate, such as silicon or fused silica, to make channels 5-10 &mu;m wide and 10 &mu;m deep (See Figure 1). The smallest dimensions, at the entrance to the mixing region, are on the order of 1 &mu;m in size. The chip is sealed with a thin glass or fused silica coverslip for optical access. Typical total linear flow rates are ~1 m/s, yielding Re~10, but the protein consumption is only ~0.5 nL/s or 1.8 &mu;L/hr. Protein concentration depends on the detection method: For tryptophan fluorescence the typical concentration is 100 &mu;M (for 1 Trp/protein) and for FRET the typical concentration is ~100 nM.
The folding process is initiated by rapid dilution of denaturant from 6 M to 0.06 M guanidine hydrochloride. The protein in high denaturant flows down a central channel and is met on either side at the mixing region by buffer without denaturant moving ~100 times faster (see Figure 2). This geometry causes rapid constriction of the protein flow into a narrow jet ~100 nm wide. Diffusion of the light denaturant molecules is very rapid, while diffusion of the heavy protein molecules is much slower, diffusing less than 1 &mu;m in 1 ms. The difference in diffusion constant of the denaturant and the protein results in rapid dilution of the denaturant from the protein stream, reducing the effective concentration of the denaturant around the protein. The protein jet flows at a constant rate down the observation channel and fluorescence of the protein during folding can be observed using a scanning confocal microscope5.

In this work we explain the fabrication and use of a microfluidic mixer capable of mixing two solutions in ~8 &mu;s. We also demonstrate the use of these mixers with spectroscopic detection using UV fluorescence and fluorescence resonance energy transfer (FRET).

The process by which a protein folds into its native conformation is highly relevant to biology and human health yet still poorly understood. One reason ...

Production, Characterization and Potential Uses of a 3D Tissue-engineered Human Esophageal Mucosal Model

The incidence of both esophageal adenocarcinoma and its precursor, Barrett&#8217;s Metaplasia, are rising rapidly in the western world. Furthermore esophageal adenocarcinoma generally has a poor prognosis, with little improvement in survival rates in recent years. These are difficult conditions to study and there has been a lack of suitable experimental platforms to investigate disorders of the esophageal mucosa.
A model of the human esophageal mucosa has been developed in the MacNeil laboratory which, unlike conventional 2D cell culture systems, recapitulates the cell-cell and cell-matrix interactions present in vivo and produces a mature, stratified epithelium similar to that of the normal human esophagus. Briefly, the model utilizes non-transformed normal primary human esophageal fibroblasts and epithelial cells grown within a porcine-derived acellular esophageal scaffold. Immunohistochemical characterization of this model by CK4, CK14, Ki67 and involucrin staining demonstrates appropriate recapitulation of the histology of the normal human esophageal mucosa.
This model provides a robust, biologically relevant experimental model of the human esophageal mucosa. It can easily be manipulated to investigate a number of research questions including the effectiveness of pharmacological agents and the impact of exposure to environmental factors such as alcohol, toxins, high temperature or gastro-esophageal refluxate components. The model also facilitates extended culture periods not achievable with conventional 2D cell culture, enabling, inter alia, the study of the impact of repeated exposure of a mature epithelium to the agent of interest for up to 20 days. Furthermore, a variety of cell lines, such as those derived from esophageal tumors or Barrett&#8217;s Metaplasia, can be incorporated into the model to investigate processes such as tumor invasion and drug responsiveness in a more biologically relevant environment.

This manuscript describes the production, characterization and potential uses of a tissue engineered 3D esophageal construct prepared from normal primary human esophageal fibroblast and squamous epithelial cells seeded within a de-cellularized porcine scaffold. The results demonstrate the formation of a mature stratified epithelium similar to the normal human esophagus.

The incidence of both esophageal adenocarcinoma and its precursor, Barrett&#8217;s Metaplasia, are rising rapidly in the western world. Furthermore ...

3D Hydrogel Scaffolds for Articular Chondrocyte Culture and Cartilage Generation

Human articular cartilage is highly susceptible to damage and has limited self-repair and regeneration potential. Cell-based strategies to engineer cartilage tissue offer a promising solution to repair articular cartilage. To select the optimal cell source for tissue repair, it is important to develop an appropriate culture platform to systematically examine the biological and biomechanical differences in the tissue-engineered cartilage by different cell sources. Here we applied a three-dimensional (3D) biomimetic hydrogel culture platform to systematically examine cartilage regeneration potential of juvenile, adult, and osteoarthritic (OA) chondrocytes. The 3D biomimetic hydrogel consisted of synthetic component poly(ethylene glycol) and bioactive component chondroitin sulfate, which provides a physiologically relevant microenvironment for in vitro culture of chondrocytes. In addition, the scaffold may be potentially used for cell delivery for cartilage repair in vivo. Cartilage tissue engineered in the scaffold can be evaluated using quantitative gene expression, immunofluorescence staining, biochemical assays, and mechanical testing. Utilizing these outcomes, we were able to characterize the differential regenerative potential of chondrocytes of varying age, both at the gene expression level and in the biochemical and biomechanical properties of the engineered cartilage tissue. The 3D culture model could be applied to investigate the molecular and functional differences among chondrocytes and progenitor cells from different stages of normal or aberrant development.

Cartilage repair represents an unmet medical challenge and cell-based approaches to engineer human articular cartilage are a promising solution. Here, we describe three-dimensional (3D) biomimetic hydrogels as an ideal tool for the expansion and maturation of human articular chondrocytes.

Human articular cartilage is highly susceptible to damage and has limited self-repair and regeneration potential. Cell-based strategies to engineer ...

A Hormone-responsive 3D Culture Model of the Human Mammary Gland Epithelium

The process of mammary epithelial morphogenesis is influenced by hormones. The study of hormone action on the breast epithelium using 2D cultures is limited to cell proliferation and gene expression endpoints. However, in the organism, mammary morphogenesis occurs in a 3D environment. 3D culture systems help bridge the gap between monolayer cell culture (2D) and the complexity of the organism. Herein, we describe a 3D culture model of the human breast epithelium that is suitable to study hormone action. It uses the commercially available hormone-responsive human breast epithelial cell line, T47D, and rat tail collagen type 1 as a matrix. This 3D culture model responds to the main mammotropic hormones: estradiol, progestins and prolactin. The influence of these hormones on epithelial morphogenesis can be observed after 1- or 2-week treatment according to the endpoint. The 3D cultures can be harvested for analysis of epithelial morphogenesis, cell proliferation and gene expression.

We describe a 3D culture model of the human breast epithelium that is suitable to study hormone action.

The process of mammary epithelial morphogenesis is influenced by hormones. The study of hormone action on the breast epithelium using 2D cultures is ...

Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung tissue from porcine, rat, or mouse, the tissue is perfused and submerged in subsequent chemical detergents to remove the cellular debris. Histological comparison of the tissue before and after processing confirms removal of over 95% of double stranded DNA and alpha galactosidase staining suggests the majority of cellular debris is removed. After decellularization, the tissue is lyophilized and then cryomilled into a powder. The matrix powder is digested for 48 hr in an acidic pepsin digestion solution and then neutralized to form the pregel solution. Gelation of the pregel solution can be induced by incubation at 37 &#176;C and can be used immediately following neutralization or stored at 4 &#176;C for up to two weeks. Coatings can be formed using the pregel solution on a non-treated plate for cell attachment. Cells can be suspended in the pregel prior to self-assembly to achieve a 3D culture, plated on the surface of a formed gel from which the cells can migrate through the scaffold, or plated on the coatings. Alterations to the strategy presented can impact gelation temperature, strength, or protein fragment sizes. Beyond hydrogel formation, the hydrogel stiffness may be increased using genipin.

This is a method to create a 3-dimensional cell culture scaffold from pulmonary extracellular matrix. Intact lung is processed into hydrogels that can support the growth of cells in three-dimensions.

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung ...

Construction of a Multilayered Mesenchymal Stem Cell Sheet with a 3D Dynamic Culture System

Stem cell therapy shows a promising future in regenerating injured organ and tissues, and the cell sheet technique has been developed to improve the low cell retention and poor survival within the target zone. However, during the in vitro construction process, a solution for maintaining stem cell bioactivity and increasing the cell amount within the cell sheet is urgently needed. Here, this protocol presents a method for constructing a multilayered cell sheet with favorable stem cell bioactivity and optimal operability. Decellularized porcine pericardium (DPP) is prepared by phospholipase A2 (PLA2) decellularization method as the cell sheet scaffold, and rat bone marrow mesenchymal stem cells (BMSCs) are isolated and expanded as the seeded cells. The temporary multilayered cell sheet structure is constructed by using RAD16-I peptide hydrogel. Finally, the cell sheet is cultured with a dynamic perfusion system to stabilize the three-dimensional (3D) structure, and the cell sheet could be obtained following a 48-hour culture in vitro. This protocol provides an efficient and feasible method for constructing a multilayered stem cell sheet, and the cell sheet could be developed as a favorable stem cell therapy product in the future.

This article provides an efficient and feasible method for constructing multilayered stem cell sheets with favorable stem cell property.

Stem cell therapy shows a promising future in regenerating injured organ and tissues, and the cell sheet technique has been developed to improve the low ...

Measuring Global Cellular Matrix Metalloproteinase and Metabolic Activity in 3D Hydrogels

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) culture systems. However, measurement of cell function in 3D culture is often more challenging. Many biological assays require retrieval of cellular material which can be difficult in 3D cultures. One way to address this challenge is to develop new materials that enable measurement of cell function within the material. Here, a method is presented for measurement of cellular matrix metalloproteinase (MMP) activity in 3D hydrogels in a 96-well format. In this system, a poly(ethylene glycol) (PEG) hydrogel is functionalized with a fluorogenic MMP cleavable sensor. Cellular MMP activity is proportional to fluorescence intensity and can be measured with a standard microplate reader. Miniaturization of this assay to a 96-well format reduced the time required for experimental set up by 50% and reagent usage by 80% per condition as compared to the previous 24-well version of the assay. This assay is also compatible with other measurements of cellular function. For example, a metabolic activity assay is demonstrated here, which can be conducted simultaneously with MMP activity measurements within the same hydrogel. The assay is demonstrated with human melanoma cells encapsulated across a range of cell seeding densities to determine the appropriate encapsulation density for the working range of the assay. After 24 h of cell encapsulation, MMP and metabolic activity readouts were proportional to cell seeding density. While the assay is demonstrated here with one fluorogenic degradable substrate, the assay and methodology could be adapted for a wide variety of hydrogel systems and other fluorescent sensors. Such an assay provides a practical, efficient and easily accessible 3D culturing platform for a wide variety of applications.

Here, a protocol is presented for encapsulating and culturing cells in poly(ethylene glycol) (PEG) hydrogels functionalized with a fluorogenic matrix metalloproteinase (MMP)-degradable peptide. Cellular MMP and metabolic activity are measured directly from the hydrogel cultures using a standard microplate reader.

Three-dimensional (3D) cell culture systems often more closely recapitulate in vivo cellular responses and functions than traditional two-dimensional (2D) ...

A Culture Method to Maintain Quiescent Human Hematopoietic Stem Cells

Human hematopoietic stem cells (HSCs), like other mammalian HSCs, maintain lifelong hematopoiesis in the bone marrow. HSCs remain quiescent in vivo, unlike more differentiated progenitors, and enter the cell cycle rapidly after chemotherapy or irradiation to treat bone marrow injury or in vitro culture. By mimicking the bone marrow microenvironment in the presence of abundant fatty acids, cholesterol, low concentration of cytokines, and hypoxia, human HSCs maintain quiescence in vitro. Here, a detailed protocol for maintaining functional HSCs in the quiescent state in vitro is described. This method will enable studies of the behavior of human HSCs under physiological conditions.

This protocol enables the maintenance of quiescent human hematopoietic stem cells in vitro by mimicking the bone marrow microenvironment using abundant fatty acids, cholesterol, lower concentrations of cytokines, and hypoxia.

Human hematopoietic stem cells (HSCs), like other mammalian HSCs, maintain lifelong hematopoiesis in the bone marrow. HSCs remain quiescent in vivo, ...