Name: measuring the rate of lipolysis in ex vivo murine adipose tissue and primary preadipocytes differentiated in vitro
Uploaded: 2023-03-17T13:00:00
Description: 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

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 wit...

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 ...

This protocol details the determination of the rate of adipocyte lipolysis in cultured adipocytes or ex vivo adipose tissue, allowing for the comparison of lipolytic rates between murine models or treatments. This protocol uses serial sampling, which allows one to internally validate that lipolysis is being measured in the linear phase and reduces measurement error. With optimization, this protocol may be used to measure lipolysis in brown adipose tissue, adipose tissue from other organisms, or adipogenic cell lines.Ex vivo tissues must be chopped into small chunks of consistent size and shape to allow for sequestration of the released free fatty acids by BSA in the media. To begin, prepare the 5%BSA medium by dissolving five grams of Bovine Serum Albumin, or BSA, in 100 milliliters of Dulbecco's Modified Eagle's Medium, or DMEM, without phenol red. Gently stir the solution to dissolve the BSA.Then prepare working concentrations of control and stimulation media. Before using, warm the media to 37 degrees Celsius. Label a 96-well plate for media collection.Inside a sterile fume hood, perform isolation and differentiation of 100%confluent primary preadipocytes. Every two to three days, change the culture media containing insulin maintaining a volume of one milliliter per well. For this study, use only cultures having differentiation rates above 90%and similar across groups.Then culture the cells in insulin free media for 24 hours before measuring lipolysis. Wash the cells with DPBS once to remove residual serum from the culture media. Prepare a 48-well plate with one well for each replicate.Place 400 microliters of room temperature DMEM into each well to be used. Use two-to-four control and two-to-four stimulated wells per tissue per mouse. Prepare a six-well plate with one well for each tissue to be collected from each mouse and put four milliliters of room temperature DMEM in each well to be used.Place the collected gonadal adipose tissue into the six-well plate. Remove tissue from the well. Place it on a silicone mat and cut it into five to seven milligram chunks with scissors.Blot the tissues on a clean towel to remove any media. Then weigh 25 to 30 milligrams, equivalent to five or six tissue chunks, for each assay well and place them in the previously prepared 48-well assay plate. Weigh the weight boat after removing the tissues and record the weight of any residue left behind.Wipe the weight boat clean between samples and retear if necessary. Use a new weight boat for each tissue. Once all the tissue samples have been weighed, place the 48-well assay plate in a 37 degree Celsius incubator under 10%carbon dioxide for 15 minutes.Perform media collection in a sterile fume hood. At time zero, after removing the media, add 400 microliters of either control or stimulation media per well and place the assay plate into an incubator at 37 degrees Celsius and 10%carbon dioxide. For ex vivo tissue culture, carefully remove media using a pipette and avoid applying suction.At times one, two, three, and four hours, collect 200 microliters of media, replace it with 200 microliters of the appropriate control or stimulation media, and return the assay plate to the incubator. Store the collection plate at four degrees Celsius. Thaw and mix the samples.Warm the reagents to room temperature and dissolve one bottle of color reagent A using one bottle of solvent A and one bottle of color reagent B using one bottle of solvent B.After creating a free fatty acid or FFA standard curve, pipette standards and samples into a 96-well assay plate. The recommended sample volume is 10 microliters per well. Add 150 microliters of reagent A to each well and mix.Incubate the assay plate at 37 degrees Celsius for five minutes. Read the absorbance of the plate at 550 nanometers and 660 nanometers and refer to this as reading A.Add 75 microliters of reagent B to each well and mix. Incubate the assay plate at 37 degrees Celsius for five minutes.Then after removing the plate from the incubator, read the absorbance of the plate again at 550 and 660 nanometers. Refer to this reading as reading B.Reconstitute the free glycerol reagent with 36 liters of ultrapure water and acclimatize it to room temperature. Thaw and mix the samples.Create a glycerol standard curve by making a 7.2 fold serial dilution of the glycerol standard solution and a water blank. Pipette 25 microliters of each standard and samples into the 96-well assay plate. Add 175 microliters of free glycerol reagent to each well and mix.Incubate the assay plate at 37 degrees Celsius for five minutes. After removing the plate from the incubator, read the absorbance of the plate at 540 nanometers. Follow the steps described in the text to calculate the lipolytic rates in the respective R square values.Then using the FFA and glycerol production rates for each sample as an individual data point, perform statistical analysis and plot the values if different lipolytic rates are being compared. If comparing ex vivo lipolytic rates 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. Then perform statistical analysis with the data.FFA and glycerol production in the in vitro differentiated preadipocytes was linear during the four-hour assay. The stimulated lipolytic rates were significantly higher than basal rates. In stimulated cells, the FFA-to-glycerol molar ratio was about three, indicating triglyceride lipolysis without significant reuptake or retention.In the unstimulated cells, the ratio was about one, suggesting a non-lipolytic source of glycerol. In ex vivo cultures, larger tissue chunks having a lower surface area to volume ratio exhibited higher FFA retention, resulting in lower FFA-to-glycerol molar ratios. In the 25 milligram tissue chunks, FFA retention negatively impacted the rates of lipolysis.Very tiny tissue chunks also exhibited reduced lipolytic rates. Reduction of BSA in the media significantly reduced both FFA and glycerol release. The FFA accumulation in the 5%BSA media after 24 hours stimulation was five millimolar while that of the 0.5%BSA media was only 0.6 millimolar.The FFA levels in the collected media reached the danger zone of 2.3 millimolar in three hours. The lack of increase in media FFA and glycerol at four hours suggested feedback inhibition. Excluding the four-hour time point caused a small, but significant increase in the release rates.The rate of FFA release was linear even between the usual time points. When removing media from the ex vivo lipolysis samples, it is important to ensure the tissue chunks are not accidentally removed by the pipette. It may be necessary to measure protein or lipid content in these samples to properly normalize the results.

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Research

JoVE Journal

Biology

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 ...

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 ...

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 ...

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 ...

Semi-Automated Isolation of Murine White Adipose Tissue

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

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

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

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

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

Quantification of Skeletal Muscle Fatty Infiltration via Decellularization

Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration

Fatty infiltration is the accumulation of adipocytes between myofibers in skeletal muscle and is a prominent feature of many myopathies, metabolic disorders, and dystrophies. Clinically in human populations, fatty infiltration is assessed using noninvasive methods, including computed tomography (CT), magnetic resonance imaging (MRI), and ultrasound (US). Although some studies have used CT or MRI to quantify fatty infiltration in mouse muscle, costs and insufficient spatial resolution remain challenging. Other small animal methods utilize histology to visualize individual adipocytes; however, this methodology suffers from sampling bias in heterogeneous pathology. This protocol describes the methodology to qualitatively view and quantitatively measure fatty infiltration comprehensively throughout intact mouse muscle and at the level of individual adipocytes using decellularization. The protocol is not limited to specific muscles or specific species and can be extended to human biopsy. Additionally, gross qualitative and quantitative assessments can be made with standard laboratory equipment for little cost, making this procedure more accessible across research laboratories.

Author Spotlight: Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration  

The present study describes decellularization-based methodologies for visualizing and quantifying intramuscular adipose tissue (IMAT) deposition through intact muscle volume, as well as quantifying metrics of individual adipocytes that comprise IMAT.

Fatty infiltration is the accumulation of adipocytes between myofibers in skeletal muscle and is a prominent feature of many myopathies, metabolic ...

Culturing Preadipocytes Isolated from Murine Perivascular Adipose Tissue

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

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

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

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

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

Murine Adipose Tissue Excision

Shave the abdominal and inguinal regions of two sedated adult male Sprague Dawley rats. Disinfect the shaved regions with an iodine solution. Make a medial longitudinal incision from the sternum to the testicular region, Using a pair of sterile forceps, remove the adipose tissue from the epididymidis and the kidneys.Place the tissue in conical tubes containing 20 milliliters of cold, sterile PBS-AA solution.