This work was supported by the US National Institutes of Health grant R01DK126944 to S.M.R.

The use of all animals was approved by the Institutional Animal Care and Use Committee (IACUC) at Weill Cornell Medical College of Cornell University.
1. Preparation of buffers and collection plates
<ol>
	<li>Make 5% bovine serum albumin (BSA) by dissolving 5 g of BSA in 100 mL of Dulbecco&#39;s modified Eagle&#39;s medium (DMEM) without phenol red. Gently stir the BSA to dissolve (shaking is counterproductive). Once the BSA is fully dissolved, filter-sterilize the media with a 0.2 &#181;m filter. Store the BSA media at 4 &#176;C for up to 1 month.</li>
	<li>Make working concentrations of control and stimulation media. Control media: 5% BSA media with vehicle control. Stimulation media: 5% BSA media with 0.5 &#181;M CL-316,243. Make fresh stimulation media for each experiment.</li>
	<li>Warm the media to be used to 37 &#176;C. Label a 96-well plate for media collection.</li>
</ol>
2. Sample preparation
<ol>
	<li>Perform cell culture as described below. Undertake all cell work in a sterile fume hood to minimize outside contamination.
	<ol>
		<li>Isolate and differentiate primary preadipocytes, as in73,74.
		<ol>
			<li>Plate primary preadipocytes at a high density, such as 1 x 105 cells/well in a 24-well plate in 1 mL/well culture media (15% fetal bovine serum (FBS) and 1x penicillin-streptomycin-glutamine in DMEM/F12).</li>
			<li>After the cells reach 100% confluency, differentiate with 5 &#181;M dexamethasone, 0.5 mM 3-isobutyl-1-methylxanthine, 1 &#181;g/mL insulin, and 1 &#181;M thiazolidinedione (TZD) in culture media for 3 days. Then, change to culture media with 1 &#181;g/mL insulin for at least 3 days to grow lipid droplets. Use 1 mL/well of media in the 24-well plate.</li>
			<li>Change the culture media (1 ml/well) with insulin every 2 or 3 days. The cells can be maintained in media with insulin for up to 2 weeks. Use only cultures in which differentiation rates are over 90% and are similar across groups for this assay, as reduced differentiation could be misinterpreted as a reduction in lipolytic rate.</li>
			<li>Culture the cells in insulin free media for 24 h prior to measuring lipolysis. 
			NOTE: Insulin in the media maintains lipid droplets, but also inhibits lipolysis. Incubation without insulin for 24 h allows for full lipolytic activation without a loss of lipid droplet volume. In some systems, the culture time without insulin may need to be shortened or extended.</li>
		</ol>
		</li>
		<li>Wash the cells with DPBS once to remove residual serum from the culture media. 
		NOTE: This protocol does not include serum starvation, which can activate lipolysis. Serum starvation may be employed at the researcher&#39;s discretion.</li>
	</ol>
	</li>
	<li>Perform ex vivo culture as described below.
	<ol>
		<li>Prepare a 6-well plate, with one well for each tissue to be collected from each mouse. Place 4 mL of room temperature DMEM in each well to be used. 
		NOTE: BSA in the collection media is not necessary.</li>
		<li>Prepare a 48-well plate for the lipolysis assay, with one well for each replicate. Place 400 &#181;L of room temperature DMEM in each well to be used. Use two to four control and two to four stimulated wells per tissue per mouse.</li>
		<li>Euthanize the mouse by cervical dislocation under anesthesia, with a secondary method such as bilateral pneumothorax. Here, we used a 32 g, 7-month-old female C57BL/6J mouse, fed with a 45% high fat diet for 4 months. 
		NOTE: This protocol can also be used for males, as well as other strains, diets, and ages.</li>
		<li>Spray with 70% ethanol and use scissors to make a small (~ 1 cm) lateral incision at the center of the abdominal skin, pull the skin apart by pinching either side with thumb and forefinger and fold the lower abdominal skin over to reveal the posterior subcutaneous depots. Locate and remove the inguinal lymph node and blunt dissect the inguinal adipose tissue immediately posterior to the inguinal lymph node using forceps.</li>
		<li>To collect the gonadal adipose tissue, make a lateral and a vertical incision in the peritoneum to access the peritoneal cavity. Hold the gonadal fat pad with tweezers and cut along the uterus (or epididymis for males) to remove the gonadal adipose tissue. Place the collected depots into a 6-well plate.</li>
		<li>Remove the tissue from well, place on a silicone mat, and cut into 5 to 7 mg chunks with scissors.</li>
		<li>Weigh out 25 to 30 mg (five or six chunks) for each assay well and place into a 48-well assay plate. Blot the tissue on a clean towel before weighing to remove any media. Weigh the weight boat after removal of the tissue and record the weight of any residue left behind. Wipe the weight boat clean between samples and re-tare if necessary. Use a new weight boat for each tissue.</li>
		<li>Once all the tissue samples have been weighed, place the 48-well assay plate in a 37 &#176;C, 10% CO2 incubator for 15 min.</li>
	</ol>
	</li>
</ol>
3. Lipolysis assay
<ol>
	<li>Perform media collection. Undertake transfer of the media and subsequent sample collection in a sterile fume hood to minimize potential contamination from outside sources.
	<ol>
		<li>At t = 0, remove the media and add 400 &#181;L per well of control or stimulation media, and place assay the plate into a 37 &#176;C, 10% CO2 incubator. For ex vivo tissue culture, carefully remove media using a pipette; suction should never be used. 
		NOTE: Alternatively, prepare a second plate with control and stimulation media, and transfer the tissues.</li>
		<li>At t = 1, 2, 3 and 4 h, collect 200 &#181;L of media, replace with 200 &#181;L of the appropriate control or stimulation media, and return the assay plate to the incubator. Store the collection plate at 4 &#176;C. To determine the FFA buffering capacity of the BSA media, use an additional collection at 24 h. 
		NOTE: The experiments can be stopped here, and the collected media can be stored at -20 &#176;C.</li>
	</ol>
	</li>
</ol>
4. FFA colorimetric assay
<ol>
	<li>Warm the reagents to room temperature and dissolve one bottle of color reagent A with one bottle of solvent A, and one bottle of color reagent B with one bottle of solvent B. From the date of reconstitution, these reagents are best used within 1 week. Discard 1 month after reconstitution.</li>
	<li>Thaw and mix the samples.</li>
	<li>Create an FFA standard curve. The standard solution is 1 mM. Use the following volume with the reagents for the standard curve: 25 &#181;L, 20 &#181;L, 15 &#181;L, 10 &#181;L, 10 &#181;L (1:2 dilution), 10 &#181;L (1:4 dilution), 10 &#181;L (1:8 dilution), and 10 &#181;L water for maximal range. For low FFA levels, 10 &#181;L of 1 mM, 0.8 mM, 0.6 mM, 0.4 mM, 0.2 mM, 0.1 mM, and 0.05 mM standard may be more applicable.</li>
	<li>Pipette standards and samples into a 96-well assay plate. The recommended sample volume is 10 &#181;L. Include three wells with the same volume of BSA media as the samples for background correction. 
	NOTE: If sample concentrations fall outside the range of the standard curve, repeat the assay, adjusting the sample volume to&#160;2-25 &#181;L.</li>
	<li>Add 150 &#181;L of reagent A to each well and mix. Avoid generating bubbles. Pop any bubbles with a fine gauge needle. Incubate the assay plate at 37 &#176;C for 5 min.</li>
	<li>Read the absorbance of the plate at 550 nm and 660 nm reference (Reading A).</li>
	<li>Add 75 &#181;L of reagent B to each well and mix. Avoid generating bubbles. Pop any bubbles with a fine gauge needle. Incubate the assay plate at 37 &#176;C for 5 min.</li>
	<li>Read the absorbance of the plate again at 550 nm and 660 nm reference (Reading B).</li>
</ol>
5. Glycerol colorimetric assay
<ol>
	<li>Reconstitute the free glycerol reagent with 36 mL of ultrapure water and acclimatize to room temperature. These reagents are best used within a few weeks. Discard 2 months after reconstitution.</li>
	<li>Thaw and mix the samples.</li>
	<li>Create a glycerol standard curve by making a seven point, 2-fold serial dilution of the glycerol standard solution and a water blank. 
	NOTE: The standard curve is relatively linear up to 25 &#181;L of 2.8 mM glycerol, but not linear at higher concentrations.</li>
	<li>Pipette 25 &#181;L each of standard and samples into the 96-well assay plate. Include three wells with the BSA media for background correction.</li>
	<li>Add 175 &#181;L of free glycerol reagent to each well and mix. Avoid generating bubbles. Pop any bubbles with a fine gauge needle. Incubate the assay plate at 37 &#176;C for 5 min.</li>
	<li>Read the absorbance of the plate at 540 nm.</li>
</ol>
6. Calculation of lipolytic rate
<ol>
	<li>Start with optical density (OD) values. For glycerol, use A540 OD values directly.&#160;Calculate the OD of the FFA assay according to the following formula: 
	OD = (Reading B: A550 - A660) - (Reading A: A550 - A660)</li>
	<li>Use the standard curve to calculate the FFA and glycerol levels in the collected samples. Plot the standard OD values on the y-axis, and on the x-axis, use standard concentrations relative to the sample volume (i.e., the concentration of the wells with 20 &#181;L of 1 mM FFA standard on a plate with 10 &#181;L samples is equal to 2 mM). Fit a linear trendline: 
	y = mx + b</li>
	<li>Visually inspect the standard curve and remove any points outside the linear range of the assay. Calculate sample concentrations using the equation: 
	Sample concentration: x = (OD - b)&#160;&#247; m</li>
	<li>Adjust and re-assay samples falling outside the linear assay range. To get the final sample concentration, subtract the concentration of the background wells containing only BSA media from the concentration of the samples.</li>
	<li>Calculate the moles of FFA and glycerol produced by each sample at each time point, according to the formula: 
	<img alt="Equation 1" src="/files/ftp_upload/65106/65106eq01.jpg" style="margin: auto;" /> 
	where Cn = concentration at time t = n; Vt = total volume in the well; Vs = sample collection volume; and Mn = moles produced at time t = n (when concentrations are in mM and volumes are in mL the output is &#181;Mol). 
	For examples, at various time points: 
	M1 = C1&#160;&#215; Vt 
	M4 = C4&#160;&#215; Vt + (C1 + C2+ C3)Vs 
	or 
	M4 = C4&#160;&#215; Vt + C3&#160;&#215; Vs + C2&#160;&#215; Vs + C1&#160;&#215;Vs</li>
	<li>Normalize to tissue weight by dividing by the tissue weight for each sample in grams to obtain units of &#181;mol/g. For cultured cells, values are presented as &#181;mol/well. Ensure that the cell number and differentiation efficiency are comparable from well to well. 
	NOTE: Differences in proliferation or differentiation efficiency will complicate the interpretation of results and require another method of normalization (e.g., normalization to protein; see discussion).</li>
	<li>Calculate the slope of the &#181;mol/g produced (y-axis) versus time (x-axis) for each sample individually.
	<ol>
		<li>In a spreadsheet, this can be done using the =SLOPE(known_ys,known_xs) function. In a new cell, type &#34;=SLOPE&#34; (then use the cursor to highlight the sample glycerol or FFA values in &#181;mol/g, then to highlight the corresponding time values).</li>
		<li>Verify the linearity of the data. R2 values are a quick way to determine linearity of the samples. In a spreadsheet, this can be done using the =RSQ(known_ys,known_xs) function, in the same manner as described in step 6.7.1, but the initial input is =RSQ. Ensure that the R2 values are &#62; 0.98; lower values indicate deviation from linearity. This can result from a measurement/sampling error or loss of linearity.
		<ol>
			<li>Another way to test linearity is to perform a linear regression for each sample and plot the residuals. In a statistical analysis software, generate an XY table with a single Y value for each time point. Select Analyze &#62; Simple linear regression and select the box for Residual Plot before hitting OK. The residual plot will appear as a new graph.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Use the FFA and glycerol production rate (i.e., slope [(&#181;mol/g/h]) for each sample as an individual data point to perform statistical analysis, and plot values if different lipolytic conditions are being compared. If lipolytic rates are being compared across genotypes, use two or three samples per animal as technical replicates, and use the average for one data point per animal, so that the sample size is equal to the number of animals.</li>
</ol>

The authors have nothing to disclose.

Here, we provide a basic protocol for measuring the rate of lipolysis in adipocytes and ex vivo adipose tissue. To quantify lipolysis, it is important to measure lipolytic rate in the linear phase. We use a serial sampling technique, where a large fraction of media is collected and replaced with fresh media at regular intervals. This semiconservative method allows for the addition of fresh BSA with FFA buffering capacity and delays feedback inhibition, extending the duration of linear lipolysis. This experimenta...

Excess nutrients are stored in white adipose tissue in the form of triglycerides in the neutral lipid core of lipid droplets. Triglyceride stores are mobilized via lipolysis, a process by which the fatty acid side chains are sequentially cleaved by adipose tissue triglyceride lipase (ATGL), hormone-sensitive lipase (HSL), and monoglyceride lipase (MGL), resulting in the release of free fatty acids (FFAs) and the glycerol backbone1,2. Lipolysis is activated by catecholamine signaling in the adipose tissue. Sympathetic nerve terminals locally release catecholamines, which bind to &#946;-adrenergic receptors on the adipocyte plasma membrane. Upon ligand binding, these G-protein coupled receptors (GPCRs) activate adenylyl cyclase via G&#945;s. Subsequent activation of protein kinase A (PKA) by cAMP results in the upregulation of both ATGL and HSL. The phosphorylation of perilipin-1 by PKA causes the dissociation of ABHD5 (also known as CGI-58), which binds and coactivates ATGL3. PKA directly phosphorylates HSL, promoting its translocation from the cytosol to the lipid droplet, where interaction with phosphorylated perilipin-1 further promotes its lipase activity4,5,6,7. The third lipase involved in lipolysis, MGL, does not appear to be regulated by catecholamine signaling8. Importantly, triglyceride synthesis in adipocytes is mediated by the glycerol lipid synthesis pathway, which does not involve the formation of monoglycerides as an intermediate; instead, glycerol-3-phosphate acyl transferases catalyze the formation of lysophosphatidic acid, which is combined with another fatty acyl-CoA to form phosphatidic acid, and then isomerized to diglycerides before the final synthesis of triglycerides (Figure 1)9,10,11.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65106/65106fig01.jpg" /> 
Figure 1: Lipolysis and glycerol lipid synthesis pathways. Top: Lipolytic pathway; enzymes shown in red: adipose tissue triglyceride lipase (ATGL), hormone sensitive lipase (HSL), and monoglyceride lipase (MGL). Bottom: glycerol lipid synthesis pathway; enzymes shown in green: diglyceride acyltransferase (DGAT), phosphatidic acid phosphatase (PAP), lysophosphatidic acid acyltransferase (LPAT, also known as LPAATs), and glycerol-3-phosphate acyltransferase (GPAT). Lipids: triglyceride (TG), diglyceride (DG), monoglyceride (MG), free fatty acid (FFA), fatty acyl-CoA (FA-CoA), lysophosphatidic acid (LPA), and phosphatidic acid (PA). Other metabolites: inorganic phosphate (Pi) and glycerol 3-phosphate (G3P). <a href="https://www.jove.com/files/ftp_upload/65106/65106fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Extracellular adenosine is another important regulator of lipolysis, working through Gs- and Gi-coupled GPCRs to impact adenyl cyclase activity. The predominant adenosine receptor in adipocytes, ADORA1, inhibits adenylyl cyclase, and thus lipolysis through the activation of Gi12. Expressed at lower levels, and primarily in brown adipocytes, ADORA2A activates lipolysis via Gs signaling13. ADORA1 impacts both basal lipolysis and the response to adrenergic agonists. The effect of adenosine on lipolysis can be controlled by adding adenosine deaminase to neutralize adenosine, as well as the ADORA1-specific agonist phenylisopropyladenosine14,15. Hormonal activation of Gq-coupled GPCRs can also affect lipolysis via the activation of phospholipase C and protein kinase C16,17,18,19. Inflammatory signals also impact lipolytic rates. TLR4 activation by LPS (and other endotoxins) increases the lipolytic rate by activating ERK, which phosphorylates perilipin-1 and HSL20. TNF-&#945; also activates lipolysis via ERK and NF-&#954;B activation, as well as transcriptional downregulation of the phosphodiesterase PDE-3B and CIDEC21,22,23. IL-6 has also been associated with increased adipocyte lipolysis, especially in mesenteric adipose tissue, whose FFA release impacts hepatic steatosis and gluconeogenesis24,25,26.
Lipolysis is suppressed during the fed state by insulin. AKT phosphorylates and activates PDE-3B to suppress cAMP signaling and prevent PKA activation27. Insulin also transcriptionally downregulates ATGL28. Obesity promotes catecholamine resistance through a variety of mechanisms, including the downregulation of &#946;-adrenergic receptors in adipocytes29,30,31,32,33. Adipocytes express all three &#946;-adrenergic receptors (&#946;-1, &#946;-2, and &#946;-3). While &#946;-1 and &#946;-2 adrenergic receptors are ubiquitously expressed, the &#946;-3 adrenergic receptor is predominately expressed in adipocytes in mice34,35. Adrb3 expression is induced by C/EBP&#945; during adipogenesis36. The &#946;-3 adrenergic receptor is highly expressed in mature adipocytes. The activation of &#946;-1 and &#946;-2 adrenergic receptors is self-limiting due to feedback inhibition by &#946;-arrestin37. Feedback inhibition of the &#946;-3 adrenergic receptor is mediated by other signaling pathways, which reduce Adrb3 expression33,38,39.
Numerous compounds can be used to activate adipocyte lipolysis. Catecholamines are major physiological activators of lipolysis. Norepinephrine (or noradrenaline) and epinephrine (or adrenaline) activate all three &#946;-adrenergic receptors40. Norepinephrine and epinephrine also effect lipolysis via activation of &#945;-adrenergic receptor signaling41. Commonly used &#946;-adrenergic receptor agonists include isoproterenol, which is a non-selective &#946;-adrenergic receptor agonist, and the &#946;-3 adrenergic receptor agonists CL-316,243 and mirabegron42. Given that adipocytes predominantly express the &#946;-3 adrenergic receptor, we use CL-316,243 as an example here. Its specificity for the &#946;-3 adrenergic receptor also makes it a relatively specific activator of adipocyte catecholamine signaling, that can also be safely used in vivo. Note that the commonly used concentration of 10 &#181;M CL-316,243 in cell culture is orders of magnitude higher than the ~0.1 &#181;M dose required to achieve a maximal response33. Forskolin bypasses the adrenergic receptor, directly activating adenylyl cyclase and downstream lipolytic signaling. There are many more activators, as well as suppressors of lipolysis. When selecting a compound to stimulate lipolysis, the receptor-specificity and downstream signaling pathways should be carefully considered within the experimental design.
The rate of lipolysis in white adipose tissue is an important metabolic factor impacting cold tolerance and nutrient availability during fasting or exercise43,44,45,46. The purpose of this protocol is to measure the rate of lipolysis in adipocytes and adipose tissue, which will facilitate the understanding of adipocyte metabolism and how it may impact the metabolic phenotype of various murine models. To quantify the lipolytic rate, we measure the appearance of lipolytic products in the media (i.e., FFAs and glycerol). The method relies on the release of lipolytic products from the adipocyte into the media. Since white adipocytes express low levels of glycerol kinase, glycerol reuptake rates are low47. Conversely, the production of FFAs and glycerol by metabolic pathways other than lipolysis should also be considered. Adipocytes appear to express a phosphatase with activity against glycerol-3 phosphate, enabling the production of glycerol from glycerol-3-phosphate derived from glucose48,49,50. Glycolysis is a source of glycerol-3-phosphate used for FFA re-esterification in white adipocytes. When glucose levels are limited, glyceroneogenesis requires other 3-carbon sources, such as lactate and pyruvate51. The channeling of FFAs released by lipolysis within the cell and their metabolic fate is poorly understood; FFAs released by lipolysis must be converted to fatty acyl-CoA, before being re-esterified or undergoing &#946;-oxidation. It appears that FFAs released by lipolysis likely exit the cell before being taken back up and converted to fatty acyl-CoA52,53,54,55,56,57,58,59,60,61,62. FFAs can be sequestered outside of the cell by albumin. Importantly, long-chain FFAs are known to feedback-inhibit lipolysis if they are not sequestered by albumin63,64,65,66,67. Thus, optimizing the FFA buffering capacity of the media during the lipolysis assay is critical. The procedure described here is similar to previously published methods to measure the lipolytic rate in adipocytes and ex vivo adipose tissue from mice and humans15,68,69,70,71. This protocol differs through the use of serial sampling; by performing serial sampling, we can internally validate that lipolysis is being measured in the linear phase and utilize multiple measurements to calculate the rate of lipolysis, thereby reducing measurement error to increase confidence in the final calculated value. The drawback of serial sampling is that the assay requires more time and reagents; however, the longer timeframe reduces impact of measurement error on the standard error of the estimates of the rate. Additionally, this protocol measures both FFA and glycerol release, and considers the ratio of FFA:glycerol release with the goal of achieving a 3:1 ratio, as would be expected from complete lipolysis and release of lipolytic products into the media72.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>24-Well tissue culture treated plate</td>
			<td>Corning Inc</td>
			<td>3527</td>
			<td>Must be tissue culture treated for adipocyte differntiation</td>
		</tr>
		<tr>
			<td>48-Well flat bottom plate with lid</td>
			<td>Corning Inc</td>
			<td>353078</td>
			<td>Can be tissue culture treated</td>
		</tr>
		<tr>
			<td>6-Well flat bottom plate with lid</td>
			<td>Corning Inc</td>
			<td>353046</td>
			<td>Can be tissue culture treated</td>
		</tr>
		<tr>
			<td>96-Well PCR Plate</td>
			<td>USA sceintific</td>
			<td>1402-9100</td>
			<td>Any conical 0.2 mL PCR plate will be convenient&#160;</td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin</td>
			<td>Sigma Aldrich</td>
			<td>A9418</td>
			<td>FFA free BSA such as A8806, is also commonly used. The BSA should not have detectable FFA, also lot to lot variations in BSA can impact the observed rate of lipolysis</td>
		</tr>
		<tr>
			<td>CL-316,243</td>
			<td>Sigma Aldrich</td>
			<td>C5976</td>
			<td>CAS #: 138908-40-4 availaible from other suppliers</td>
		</tr>
		<tr>
			<td>CO2 incubator</td>
			<td>PHCBI</td>
			<td>MCO-170AICUVH</td>
			<td>CO2 should ideally be set to 10% for adipose tissue, however 5% CO2 will also work</td>
		</tr>
		<tr>
			<td>DMEM, low glucose, no phenol red</td>
			<td>Thermofischer</td>
			<td>11054020</td>
			<td>Any phenol red free media should work, DMEM/F12, RPMI, but should contain volatile buffering capacity, i.e. biocarbonate</td>
		</tr>
		<tr>
			<td>FFA-free Bovine serum albumin</td>
			<td>Equitech-Bio, Inc,&#160;</td>
			<td>BAH66</td>
			<td></td>
		</tr>
		<tr>
			<td>Free Glycerol Reagent</td>
			<td>Sigma Aldrich</td>
			<td>F6428</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycerol Standard Solution</td>
			<td>Sigma Aldrich</td>
			<td>G7793&#160;</td>
			<td>This can also be made by diluting glycerol to the desired concentration</td>
		</tr>
		<tr>
			<td>HR Series NEFA Standard Solution</td>
			<td>Fujifilm</td>
			<td>276-76491</td>
			<td></td>
		</tr>
		<tr>
			<td>HR Series NEFA-HR (2) Color Reagent A</td>
			<td>Fujifilm</td>
			<td>999-34691</td>
			<td></td>
		</tr>
		<tr>
			<td>HR Series NEFA-HR (2) Color Reagent B</td>
			<td>Fujifilm</td>
			<td>991-34891</td>
			<td></td>
		</tr>
		<tr>
			<td>HR Series NEFA-HR (2) Solvent A&#160;</td>
			<td>Fujifilm</td>
			<td>995-34791</td>
			<td></td>
		</tr>
		<tr>
			<td>HR Series NEFA-HR (2) Solvent B&#160;</td>
			<td>Fujifilm</td>
			<td>993-35191</td>
			<td></td>
		</tr>
		<tr>
			<td>Microbiological Incubator</td>
			<td>Fischer Scientific</td>
			<td>S28668</td>
			<td>Any incubator at 37C can be used</td>
		</tr>
		<tr>
			<td>Nunc MicroWell 96-Well Plates</td>
			<td>Thermo Scientific</td>
			<td>269620</td>
			<td>Any optically clear, flat bottom 96-well plate works</td>
		</tr>
		<tr>
			<td>Silicone Laboratory Benchtop Mat</td>
			<td>VWR</td>
			<td>76045-300</td>
			<td>Glass plate can also be used. Absorbant surfaces are not recommended</td>
		</tr>
		<tr>
			<td>Spectrophotometer/Microplate Reader</td>
			<td>Molecular devices</td>
			<td>SpectraMax i3x&#160;</td>
			<td>Any plate reader that can read at 540, 550 and 660 mm will work</td>
		</tr>
		<tr>
			<td>V Bovine serum albumin</td>
			<td>Sigma-Aldrich</td>
			<td>810531</td>
			<td></td>
		</tr>
		<tr>
			<td>WypAll X70 Wipers</td>
			<td>Kimberly-Clark</td>
			<td>41200</td>
			<td>Any high quality paper towel will work</td>
		</tr>
	</tbody>
</table>

measuring the rate of lipolysis in ex vivo murine adipose tissue and primary preadipocytes differentiated in vitro

Adipocytes store energy in the form of triglycerides in lipid droplets. This energy can be mobilized via lipolysis, where the fatty acid side chains are sequentially cleaved from the glycerol backbone, resulting in the release of free fatty acids and glycerol. Due to the low expression of glycerol kinase in white adipocytes, glycerol re-uptake rates are negligible, while fatty acid re-uptake is dictated by the fatty acid binding capacity of media components such as albumin. Both glycerol and fatty acid release into media can be quantified by colorimetric assays to determine the lipolytic rate. By measuring these factors at multiple time points, one can determine the linear rate of lipolysis with high confidence. Here, we provide a detailed protocol for the measurement of lipolysis in in vitro differentiated adipocytes and ex vivo adipose tissue from mice. This protocol may also be optimized for other preadipocyte cell lines or adipose tissue from other organisms; considerations and optimization parameters are discussed. This protocol is designed to be useful in determining and comparing the rate of adipocyte lipolysis between mouse models and treatments.

We measured the basal and stimulated lipolytic rate of in vitro differentiated adipocytes. Primary preadipocytes from inguinal white adipose tissue were differentiated into adipocytes by the treatment of confluent cells with 5 &#181;M dexamethasone, 0.5 mM IBMX, 1 &#181;g/mL insulin, and 1 &#181;M troglitazone for 4 days, followed by an additional 3 day treatment with 1 &#181;g/mL insulin. Cells were incubated in media without insulin for 24 h prior to the lipolysis assay. At time = 0h, the cells were washed onc...

Watch this Scientific Journal Video about Measuring the Rate of Lipolysis in Ex Vivo Murine Adipose Tissue and Primary Preadipocytes Differentiated In Vitro at JoVE.com

Measuring the Rate of Lipolysis in Ex Vivo Murine Adipose Tissue and Primary Preadipocytes Differentiated In Vitro

Triglyceride lipolysis in adipocytes is an important metabolic process resulting in the liberation of free fatty acids and glycerol. Here, we provide a detailed protocol to measure basal and stimulated lipolysis in adipocytes and ex vivo adipose tissue from mice.

Measuring the Rate of Lipolysis in Ex Vivo Murine Adipose Tissue and Primary Preadipocytes Differentiated In Vitro

Adipocytes store energy in the form of triglycerides in lipid droplets. This energy can be mobilized via lipolysis, where the fatty acid side chains are ...

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JoVE Journal

Biology

2.6K Views.  Weill Cornell Medicine. Triglyceride lipolysis in adipocytes is an important metabolic process resulting in the liberation of free fatty acids and glycerol. Here, we provide a detailed protocol to measure basal and stimulated lipolysis in adipocytes and ex vivo adipose tissue from mice.

Video: Measuring the Rate of Lipolysis in Ex Vivo Murine Adipose Tissue and Primary Preadipocytes Differentiated In Vitro

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti. 

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the inner ear (fig 1). Hair cells in the developing cochlea, which are the mechanosensory cells of the auditory system, are aligned in one row of inner hair cells and three (in the base and mid-turns) to four (in the apical turn) rows of outer hair cells that span the length of the organ of Corti. Hair cells transduce sound-induced mechanical vibrations of the basilar membrane into neural impulses that the brain can interpret. Most cases of sensorineural hearing loss are caused by death or dysfunction of cochlear hair cells.
An increasingly essential tool in auditory research is the isolation and in vitro culture of the organ explant 1,2,9. Once isolated, the explants may be utilized in several ways to provide information regarding normative, anomalous, or therapeutic physiology. Gene expression, stereocilia motility, cell and molecular biology, as well as biological approaches for hair cell regeneration are examples of experimental applications of organ of Corti explants.
This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

This procedure describes a method for the isolation and culture of the murine organ of Corti with or without the spiral limbus and spiral ganglion neurons. We also demonstrate a method for the expression of an exogenous reporter gene in the organ of Corti explant by electroporation.

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the ...

Measuring Glutathione-induced Feeding Response in Hydra

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. The movement of prey organisms causes mechanosensory discharge of the stinging cells called nematocysts from hydra, which are inserted into the prey. The feeding response in hydra, which includes curling of the tentacles to bring the prey towards the mouth, opening of the mouth and consequent engulfing of the prey, is triggered by GSH present in the fluid released from the injured prey. To be able to identify the molecular mechanism of the feeding response in hydra which is unknown to date, it is necessary to establish an assay to measure the feeding response. Here, we describe a simple method for the quantitation of the feeding response in which the distance between the apical end of the tentacle and mouth of hydra is measured and the ratio of such distance before and after the addition of GSH is determined. The ratio, called the relative tentacle spread, was found to give a measure of the feeding response. This assay was validated using a starvation model in which starved hydra show an enhanced feeding response in comparison with daily fed hydra.

Here we describe a simple assay for the quantification of the feeding response in hydra induced by the reduced form of glutathione. This assay relies on measuring the distance between the apical end of the tentacle and mouth of hydra.

Hydra is among the most primitive organisms possessing a nervous system and chemosensation for detecting reduced glutathione (GSH) for capturing the prey. ...

The c-FOS Protein Immunohistological Detection: A Useful Tool As a Marker of Central Pathways Involved in Specific Physiological Responses In Vivo and Ex Vivo

Many studies seek to identify and map the brain regions involved in specific physiological regulations. The proto-oncogene c-fos, an immediate early gene, is expressed in neurons in response to various stimuli. The protein product can be readily detected with immunohistochemical techniques leading to the use of c-FOS detection to map groups of neurons that display changes in their activity. In this article, we focused on the identification of brainstem neuronal populations involved in the ventilatory adaptation to hypoxia or hypercapnia. Two approaches were described to identify involved neuronal populations in vivo in animals and ex vivo in deafferented brainstem preparations. In vivo, animals were exposed to hypercapnic or hypoxic gas mixtures. Ex vivo, deafferented preparations were superfused with hypoxic or hypercapnic artificial cerebrospinal fluid. In both cases, either control in vivo animals or ex vivo preparations were maintained under normoxic and normocapnic conditions. The comparison of these two approaches allows the determination of the origin of the neuronal activation i.e., peripheral and/or central. In vivo and ex vivo, brainstems were collected, fixed, and sliced into sections. Once sections were prepared, immunohistochemical detection of the c-FOS protein was made in order to identify the brainstem groups of cells activated by hypoxic or hypercapnic stimulations. Labeled cells were counted in brainstem respiratory structures. In comparison to the control condition, hypoxia or hypercapnia increased the number of c-FOS labeled cells in several specific brainstem sites that are thus constitutive of the neuronal pathways involved in the adaptation of the central respiratory drive.

The c-FOS Protein Immunohistological Detection: A Useful Tool As a Marker of Central Pathways Involved in Specific Physiological Responses In Vivo and Ex Vivo

Here, we present a protocol based on c-FOS protein immunohistological detection, a classical technique used for the identification of neuronal populations involved in specific physiological responses in vivo and ex vivo.

Many studies seek to identify and map the brain regions involved in specific physiological regulations. The proto-oncogene c-fos, an immediate early gene, ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte precursors, immune cells, fibroblasts, blood vessels, and nerve projections. Although the molecular control of cell type specification and how these cells interact have been increasingly delineated, a more comprehensive understanding of these adipose-resident cells can be achieved by visualizing their distribution and architecture throughout the whole tissue. Existing immunohistochemistry and immunofluorescence approaches to analyze adipose histology rely on thin paraffin-embedded sections. However, thin sections capture only a small portion of tissue; as a result, the conclusions can be biased by what portion of tissue is analyzed. We have therefore developed an adipose tissue clearing technique, Adipo-Clear, to permit comprehensive three-dimensional visualization of molecular and cellular patterns in whole adipose tissues. Adipo-Clear was adapted from iDISCO/iDISCO+, with specific modifications made to completely remove the lipid stored in the tissue while preserving native tissue morphology. In combination with light-sheet fluorescence microscopy, we demonstrate here the use of the Adipo-Clear method to obtain high-resolution volumetric images&#160;of an entire adipose tissue.

Due to the high lipid content, adipose tissue has been challenging to visualize using traditional histological methods. Adipo-Clear is a tissue clearing technique that allows robust labeling and high-resolution volumetric fluorescent imaging of adipose tissue. Here, we describe the methods for sample preparation, pretreatment, staining, clearing, and mounting for imaging.

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte ...

Exploring Adipose Tissue Structure by Methylsalicylate Clearing and 3D Imaging

Obesity is a major worldwide public health issue that increases the risk to develop cardiovascular diseases, type-2 diabetes, and liver diseases. Obesity is characterized by an increase in adipose tissue (AT) mass due to adipocyte hyperplasia and/or hypertrophia, leading to profound remodeling of its three-dimensional structure. Indeed, the maximal capacity of AT to expand during obesity is pivotal to the development of obesity-associated pathologies. This AT expansion is an important homeostatic mechanism to enable adaptation to an excess of energy intake and to avoid deleterious lipid spillover to other metabolic organs, such as muscle and liver. Therefore, understanding the structural remodeling that leads to the failure of AT expansion is a fundamental question with high clinical applicability. In this article, we describe a simple and fast clearing method that is routinely used in our laboratory to explore the morphology of mouse and human white adipose tissue by fluorescent imaging. This optimized AT clearing method is easily performed in any standard laboratory equipped with a chemical hood, a temperature-controlled orbital shaker and a fluorescent microscope. Moreover, the chemical compounds used are readily available. Importantly, this method allows one to resolve the 3D AT structure by staining various markers to specifically visualize the adipocytes, the neuronal and vascular networks, and the innate and adaptive immune cells distribution.

Here, we describe a simple, inexpensive and fast clearing method to resolve the 3D structure of both mouse and human white adipose tissue using a combination of markers to visualize vasculature, nuclei, immune cells, neurons, and lipid-droplet coat proteins by fluorescent imaging.

Obesity is a major worldwide public health issue that increases the risk to develop cardiovascular diseases, type-2 diabetes, and liver diseases. Obesity ...

Isolation and Differentiation of Primary White and Brown Preadipocytes from Newborn Mice

The understanding of the mechanisms underlying adipocyte differentiation and function has greatly benefited from the use of immortalized white preadipocyte cell lines. These cultured cell lines, however, have limitations. They do not fully capture the diverse functional spectrum of the heterogenous adipocyte populations that are now known to exist within white adipose depots. To provide a more physiologically relevant model to study the complexity of white adipose tissue, a protocol has been developed and optimized to enable simultaneous isolation of primary white and brown adipocyte progenitors from newborn mice, their rapid expansion in culture, and their differentiation in vitro into mature, fully functional adipocytes. The primary advantage of isolating primary cells from newborn, rather than adult mice, is that the adipose depots are actively developing and are, therefore, a rich source of proliferating preadipocytes. Primary preadipocytes isolated using this protocol differentiate rapidly upon reaching confluence and become fully mature in 4-5 days, a temporal window that accurately reflects the appearance of developed fat pads in newborn mice. Primary cultures prepared using this strategy can be expanded and studied with high reproducibility, making them suitable for genetic and phenotypic screens and enabling the study of the cell-autonomous adipocyte phenotypes of genetic mouse models. This protocol offers a simple, rapid, and inexpensive approach to study the complexity of adipose tissue in vitro.

This report describes a protocol for the simultaneous isolation of primary brown and white preadipocytes from newborn mice. Isolated cells can be grown in culture and induced to differentiate into fully mature white and brown adipocytes. The method enables genetic, molecular, and functional characterization of primary fat cells in culture.

The understanding of the mechanisms underlying adipocyte differentiation and function has greatly benefited from the use of immortalized white ...

Immunophenotyping and Cell Sorting of Human MKs from Human Primary Sources or Differentiated In Vitro from Hematopoietic Progenitors

Megakaryocyte (MK) differentiation encompasses a number of endomitotic cycles that result in a highly polyploid (reaching even&#160;&#62;64N) and extremely large cell (40-60 &#181;m). As opposed to the fast-increasing knowledge in megakaryopoiesis at the cell biology and molecular level, the characterization of megakaryopoiesis by flow cytometry is limited to the identification of mature MKs using lineage-specific surface markers, while earlier MK differentiation stages remain unexplored. Here, we present an immunophenotyping strategy that allows the identification of successive MK differentiation stages, with increasing ploidy status, in human primary sources or in vitro cultures with a panel integrating MK specific and non-specific surface markers. Despite its size and fragility, MKs can be immunophenotyped using the above-mentioned panel and enriched by fluorescence-activated cell sorting under specific conditions of pressure and nozzle diameter. This approach facilitates multi-Omics studies, with the aim to better understand the complexity of megakaryopoiesis and platelet production in humans. A better characterization of megakaryopoiesis may pose fundamental in the diagnosis or prognosis of lineage-related pathologies and malignancy.

Immunophenotyping and Cell Sorting of Human MKs from Human Primary Sources or Differentiated In Vitro from Hematopoietic Progenitors

Here, we present an immunophenotyping strategy for the characterization of megakaryocyte differentiation, and show how that strategy allows the sorting of megakaryocytes at different stages with a fluorescence-activated cell sorter. The methodology can be applied to human primary tissues, but also to megakaryocytes generated in culture in vitro.

Megakaryocyte (MK) differentiation encompasses a number of endomitotic cycles that result in a highly polyploid (reaching even&#160;&#62;64N) and ...

Differentiated Mouse Adipocytes in Primary Culture: A Model of Insulin Resistance

Insulin resistance is a reduced effect of insulin on its target cells, usually derived from decreased insulin receptor signaling. Insulin resistance contributes to the development of type 2 diabetes (T2D) and other obesity-derived diseases of high prevalence worldwide. Therefore, understanding the mechanisms underlying insulin resistance is of great relevance. Several models have been used to study insulin resistance both in vivo and in vitro; primary adipocytes represent an attractive option to study the mechanisms of insulin resistance and identify molecules that counteract this condition and the molecular targets of insulin-sensitizing drugs. Here, we have established an insulin resistance model using primary adipocytes in culture treated with tumor necrosis factor-&#945; (TNF-&#945;).
Adipocyte precursor cells (APCs), isolated from collagenase-digested mouse subcutaneous adipose tissue by magnetic cell separation technology, are differentiated into primary adipocytes. Insulin resistance is then induced by treatment with TNF-&#945;, a proinflammatory cytokine that reduces the tyrosine phosphorylation/activation of members of the insulin signaling cascade. Decreased phosphorylation of insulin receptor (IR), insulin receptor substrate (IRS-1), and protein kinase B (AKT) are quantified by western blot. This method provides an excellent tool to study the mechanisms mediating insulin resistance in adipose tissue.

This protocol describes the isolation of mouse preadipocytes from subcutaneous fat, their differentiation into mature adipocytes, and the induction of insulin resistance. Insulin action is evaluated by the phosphorylation/activation of members of the insulin signaling pathway through western blot. This method allows direct determination of insulin resistance/sensitivity in primary adipocytes.

Insulin resistance is a reduced effect of insulin on its target cells, usually derived from decreased insulin receptor signaling. Insulin resistance ...

Adipose Tissue-Derived Stem Cell Isolation and Characterization

Isolation and Identification of Mesenchymal Stem Cells Derived from Adipose Tissue of Sprague Dawley Rats

Adult mesenchymal cells have revolutionized molecular and cell biology in recent decades. They can differentiate into different specialized cell types, in addition to their great capacity for self-renewal, migration, and proliferation. Adipose tissue is one of the least invasive and most accessible sources of mesenchymal cells. It has also been reported to have higher yields compared to other sources, as well as superior immunomodulatory properties. Recently, different procedures for obtaining adult mesenchymal cells from different tissue sources and animal species have been published. After evaluating the criteria of some authors, we standardized a methodology that is applicable to different purposes and easily reproducible. A pool of stromal vascular fraction (SVF) from perirenal and epididymal adipose tissue allowed us to develop primary cultures with optimal morphology and functionality. The cells were observed adhered to the plastic surface for 24 h, and exhibited a fibroblast-like morphology, with prolongations and a tendency to form colonies. Flow cytometry (FC) and immunofluorescence (IF) techniques were used to assess the expression of the membrane markers CD105, CD9, CD63, CD31, and CD34. The ability of adipose-derived stem cells (ASCs) to differentiate into the adipogenic lineage was also assessed using a cocktail of factors (4 &#181;M insulin, 0.5 mM 3-methyl-iso-butyl-xanthine, and 1 &#181;M dexamethasone). After 48 h, a gradual loss of fibroblastoid morphology was observed, and at 12 days, the presence of lipid droplets positive to oil red staining was confirmed. In summary, a procedure is proposed to obtain optimal and functional ASC cultures for application in regenerative medicine.

Author Spotlight: Isolation and Identification of Mesenchymal Stem Cells Derived from Adipose Tissue of Sprague Dawley Rats  

This protocol describes a methodology for isolating and identifying adipose tissue-derived mesenchymal stem cells (MSCs) from Sprague Dawley rats.

Adult mesenchymal cells have revolutionized molecular and cell biology in recent decades. They can differentiate into different specialized cell types, in ...