We thank the Sanchez-Gurmaches and Wallace lab members for valuable discussions. This work was supported by grants from the American Heart Association (18CDA34080527 to JSG and 19POST34380545 to RM), the NIH (R21OD031907 to JSG), a CCHMC Trustee Award, a CCHMC Center for Pediatric Genomics Award, and a CCHMC Center for Mendelian Genomics &#38; Therapeutics Award. This work was supported in part by NIH P30 DK078392 of the Digestive Diseases Research Core Center in Cincinnati. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. RT and MW were supported by a UCD Ad Astra Fellowship.

All experiments were approved by the Institutional Animal Care and Use Committee at Cincinnati Children&#39;s Hospital Medical Center.
1. Preparation of D2O
NOTE: To avoid experimental variation, prepare sufficient solution/drinking water for all mice for the duration of the experiment.
<ol>
	<li>For intraperitoneal injection: generate 0.9% w/v saline D2O by dissolving 9 g of NaCl per liter of D2O. Filter through a non-pyrogenic 0.2 &#181;m filter to sterilize.</li>
	<li>For drinking D2O-water: generate 8% v/v D2O-enriched water to be used as drinking water by mixing 80 mL of D2O per 920 mL of regular drinking water. Regular drinking water can be obtained from the mouse facility. Filter through a non-pyrogenic 0.2 &#181;m filter to sterilize.</li>
</ol>
2. Modulation of BAT activity by temperature acclimation
<ol>
	<li>Separate the mice so there are two mice per cage 2 weeks prior to the start of temperature acclimation. Change the water supply to water bottles for the mice to adapt to and maintain all the cages at 22 &#176;C.</li>
	<li>Prepare the mouse environmental chambers 1 week before starting the temperature acclimation by setting the appropriate temperatures: 30 &#176;C for thermoneutrality, 22 &#176;C for room temperature, and 18 &#176;C for cold exposure.</li>
	<li>At the start of temperature acclimation, replace the cages with new ones with no environmental enrichment (to avoid nests). Move the cages to their respective temperatures. 
	NOTE: Cages assigned to thermoneutrality will stay at 30 &#176;C for 4 weeks. Cages assigned to room temperature will stay at 22 &#176;C for 4 weeks. Cages assigned to cold exposure will become progressively colder on a weekly schedule: 18 &#176;C for the first week, 14 &#176;C for the second week, 10 &#176;C for the third week, and 6 &#176;C for the fourth week.</li>
	<li>Change soiled cages weekly for all conditions. Additionally, replace food and water bottles with new ones preadapted at the appropriate temperature for at least 24 h. 
	NOTE: Be mindful about the food amounts added every week, especially for mice in the cold, since they will consume a significantly greater amount of food compared to normal conditions. Provide food ad libitum.</li>
</ol>
3. Administration of D&#8203;2O
<ol>
	<li>Inject each animal with 0.9% w/v saline-D2O at 35 &#181;L/g of body weight, 12 h-3 days before tissue collection, via intraperitoneal injection using a 1 mL syringe and 26 G needle. 
	NOTE: Please see the discussion section for more information on selecting an appropriate labelling time.</li>
	<li>Change the water bottles to bottles containing 8% v/v D2O-enriched drinking water.</li>
</ol>
4. Plasma and tissue collection, processing, and storage
<ol>
	<li>At the end of the experiment, sacrifice the mice using approved methodologies (e.g., carbon dioxide overdose followed by cervical dislocation).
	<ol>
		<li>Use heat/cool pads or other methods to avoid sudden temperature changes before euthanasia that may affect the results. Euthanize the mice following approved methodologies. Proceed immediately to blood and tissue collection.</li>
	</ol>
	</li>
	<li>Collect blood through cardiac puncture using a 26 G needle and store in an ethylenediaminetetraacetic acid blood collection tube. Keep the blood on ice until further processing.</li>
	<li>Cut open the skin along the middle line of the back of the mouse from the lower area of the thoracic cavity up to the upper area of the neck, while pulling the skin up to avoid affecting tissues below the skin. The interscapular BAT is located between the shoulder blades under a thin layer of white adipose tissue, and it is composed of two pyramidal shaped lobes.
	<ol>
		<li>Clean the tissue by washing in ice cold phosphate buffered saline (PBS). Dab on a paper towel to eliminate excess liquid, weigh on an analytical scale, and collect in a microtube.</li>
		<li>Immediately flash freeze the tissue using liquid nitrogen. Other BAT depots can also be collected.</li>
	</ol>
	</li>
	<li>Centrifuge the blood samples at 10,000 x g for 10 min at 4 &#176;C. After centrifugation, carefully collect the plasma without disturbing the red blood cell pellet. Transfer it to a new ice-cold microtube and flash freeze in liquid nitrogen.</li>
	<li>Store brown fat and plasma samples at -80 &#176;C until use.</li>
</ol>
5. Lipid extraction from adipose tissue
<ol>
	<li>Before beginning the extraction
	<ol>
		<li>Prepare a 1 mM solution of hexadecenoic-d31 acid in methanol in a glass vial. This will serve as the internal fatty acid standard.</li>
		<li>Pre-cool the required amount of chloroform (CHCl3) and methanol (CH3OH) in a -80 &#176;C freezer, or on dry ice. 
		NOTE: The antioxidant di-tert-butyl-4-methylphenol (BHT) may be added to the CHCl3 at a concentration of 0.01% w/v (2.5 mg/25 mL) to prevent oxidation of the double bonds in unsaturated fatty acids.</li>
		<li>Pre-label microcentrifuge tubes for each tissue sample and an extra tube to be used for a blank extraction. 
		CAUTION: CH3OH and CHCl3 are highly volatile and toxic if inhaled. Use only in fume hoods. 
		NOTE: Different brands of microcentrifuge tubes have differing levels of background palmitate and capabilities in terms of preventing solvent leakage. We recommend that a variety of tubes be tested first, to ensure that solvent leakage is prevented and that the tubes have minimal levels of contaminating palmitate. Please see Yao et al.32 for further discussion.</li>
	</ol>
	</li>
	<li>Take the samples out of the freezer and place on dry ice.</li>
	<li>Place the pre-labelled microcentrifuge tube on an analytical balance and tare the balance. Place tweezers and a scalpel/steel razor blade on dry ice for 10-20 s to cool.
	<ol>
		<li>Use the tweezers to take the frozen tissue sample out of the tube and place on a plastic weigh boat. The weigh boat may be placed on a flat block of dry ice or another pre-cooled surface.</li>
		<li>Using the scalpel or steel razor blade, dissect a small portion of the tissue, equivalent to 5-15 mg in weight. Place in the microcentrifuge tube and record the exact weight. Repeat for each specimen. Samples can be stored in the freezer at this point or can be advanced to the steps below for lipid extraction. 
		NOTE: Ensure that the scalpel is properly cleaned with 70% ethanol between samples and a fresh weigh boat is used between each sample.</li>
	</ol>
	</li>
	<li>Add 1 &#181;L/mg of 10 mM hexadecenoic-d31(C16:0-d31) acid to each sample. 
	CAUTION: The following steps (5.4 to 5.8) should be carried out under a fume hood due to the inhalation risk of the solvents.</li>
	<li>Add 250 &#181;L of CH3OH, 250 &#181;L of H2O, and 500 &#181;L of CHCl3 to each sample with three 5 mm stainless steel beads. Place the tubes in a pre-cooled block of a grinding mill and mix the samples at a vibrational frequency of 25 Hz for 5 min, or use guidelines recommended by the manufacturer for tissue samples. Remove the beads using a magnet.</li>
	<li>Centrifuge the samples at 12,000 x g for 10 min at 4 &#176;C. 
	NOTE: After centrifugation, a clear biphasic separation should be observed with the upper aqueous phase containing polar metabolites and the lower organic phase containing lipids and fatty acids. If no separation is seen, add 250 &#181;L of H2O and repeat the vortex and centrifugation steps.</li>
	<li>Using a micropipette, take a fixed volume from the bottom phase of each sample into correspondingly labelled microcentrifuge tubes.</li>
	<li>Add 500 &#181;L of CHCl3 to the remaining sample and repeat steps 5.6-5.7.</li>
	<li>Place the samples under nitrogen gas or in a CHCl3-resistant refrigerated centrifugal vacuum at 4 &#176;C until completely dry. Dried samples can be stored at -20 &#176;C until ready for derivatization. 
	NOTE: The top layer of each sample can also be collected at this point and dried as in step 5.9 in order to analyze polar metabolites.</li>
</ol>
6. Preparation of fatty acid methyl esters (FAMEs) and GCMS analysis
<ol>
	<li>Acid-catalyzed esterification and transesterification to prepare FAMEs 
	CAUTION: The following steps should be carried out under a fume hood due to the inhalation risk of the solvents.
	<ol>
		<li>If the samples have been stored in freezer, dry under nitrogen for 5 min to ensure that no water is present.</li>
		<li>Using MS-grade solvents, pipette 98 mL of anhydrous CH3OH into a glass media bottle. Slowly add 2 mL of anhydrous sulfuric acid in the fume hood to make 2% H2SO4 in CH3OH. Mix by swirling the closed bottle.</li>
		<li>Add 500 &#181;L of 2% H2SO4 in CH3OH solution to each sample and vortex briefly.</li>
		<li>Incubate the samples on a heat block at 50 &#176;C for 2 h.</li>
		<li>Remove the samples from the heat block and add 100 &#181;L of saturated NaCl solution and 500 &#181;L of hexane to each sample.</li>
		<li>Vortex the samples vigorously at room temperature for 1 min. Leave the samples to sit for 1 min; two phases should be apparent after this.</li>
		<li>Collect the upper phase into a fresh microcentrifuge tube (see note in section 5.1.3 for appropriate selection of the microcentrifuge tube).</li>
		<li>To maximize the yield, repeat steps 6.1.5-6.1.7, collecting the second sample into the same labelled tubes.</li>
		<li>Dry the samples at room temperature under nitrogen gas.</li>
		<li>Resuspend the samples in 20 &#181;L/mg of hexane, relative to the original tissue weight, and transfer immediately to glass GC vial with a glass insert. 
		NOTE: Work quickly while transferring samples to minimize evaporation.</li>
	</ol>
	</li>
	<li>GCMS analysis
	<ol>
		<li>To determine the abundance of FAME isotopologues, inject the samples on a single quadrupole gas chromatography mass spectrometer (GCMS). 
		NOTE: While many column types can be used to detect palmitate, the following temperature program was established for a GCMS column that has been developed for the separation of fatty acid cis/trans isomers, as detailed in the Table of Materials. This column has a length of 50 m with an 0.25 mm inner diameter.</li>
		<li>Inject 1 &#181;L of sample in a split/splitless inlet at an inlet temperature of 270 &#176;C, using helium as the carrier gas, flowing at 1 mL/min. Use a splitless injection for low abundant fatty acids with a total flow of 19 mL/min, a septum purge of 3 mL/min, and a purge flow to split vent of 15 mL/min at 0.75 min. Use a split ratio of 10:1-40:1 for high abundant fatty acids such as palmitate and oleate.</li>
		<li>Use the following oven parameters: an initial temperature of 80 &#176;C; increase by 20 &#176;C/min increments to 170 &#176;C; increase by 1 &#176;C/min increments to 204 &#176;C; increase by 20 &#176;C/min increments to 250 &#176;C; and then hold at 250 &#176;C for 10 min.</li>
		<li>Use the following mass selective detector (MSD) parameters: an electron impact ionization mode of 70 eV and scan over the range of 50-400 m/z with a scan speed of 1,562 (u/s) and a frequency of 4.1 scans/s. Use a transfer line at 280 &#176;C, an ion source at 230 &#176;C, and a quadrupole at 150 &#176;C. 
		&#8203;NOTE: Other columns may be used, but the temperature program will vary.</li>
		<li>Sample sequence: Randomize the sample order of injection and inject at least two or three hexane blanks at the beginning of the sequence, after every fifth sample within the sequence, and two or three at the end of the sequence.</li>
		<li>Use instrument specific software, or free open-access software such as Metabolite-Detector33, to integrate the ions in Table 1 for each fatty acid methyl ester. 
		NOTE: We have outlined ions that cover M1-M5 isotopologues in Table 1 that we have found covers the amount of deuterium incorporation with this labelling window. However, this may need to be expanded if de novo flux is significantly higher and/or a longer labelling time is used.</li>
		<li>Use the abundance of each integrated ion to generate a mass isotopomer distribution, where the ion intensities can be converted to fractional abundance so that the sum of the mass isotopomer distribution equals one. Please see the example spreadsheet in the Supplementary File. 
		NOTE: In order to accurately determine deuterium incorporation, correction of the natural isotope abundance then has to be employed to allow for the presence of naturally occurring isotopes such as 13C, 15N, and 2H. This is performed by applying a correction matrix as outlined by Fernandez et al.34,35, and cannot be performed by subtracting the MID of an unlabelled measured metabolite from a labelled metabolite. In practice, we recommend the use of freely available software such as fluxfix36, polyMID37, or IsoCor38 to turn raw data into fractional MIDs, and we have provided the formula for isotope correction for the methyl ester products of FAS in Table 1.</li>
		<li>To determine the quantity of palmitate present, utilize the following formula: 
		<img alt="Equation 1" src="/files/ftp_upload/64219/64219eq1v2.png" style="margin: auto; height: 37px;" />&#160; 
		where ion count refers to the sum of all palmitate isotopologues integrated in Table 1 and C16:0-d31 refers to the internal standard hexadecenoic-d31. The internal standard forms a separate peak to that of the endogenous palmitate. The relative abundance of other fatty acids may also be determined, but fatty acid specific isotope internal standards (with mass shifts greater than that observed from D2O incorporation) or external standard curves may need to be employed for full quantitation. Use the blank extracted sample to determine the amount of background palmitate and subtract this from the final tissue value.</li>
		<li>Calculate the molar enrichment (ME) of palmitate by the following equation: 
		<img alt="Equation 2" src="/files/ftp_upload/64219/64219eq2v2.png" style="margin: auto; height: 65px;" /> 
		&#8203;where Mi is the normalized, fractional abundance of a palmitate isotopologue, and n is the number of possible palmitate isotopologues. For example, the ME of a palmitate molecule with the following fractional distribution, M1 = 0.25, M2 = 0.08, M3 = 0.02, is: (0.025*1) + (0.08*2) + (0.02*3) = 0.245 (See Supplementary File).</li>
	</ol>
	</li>
</ol>
7. Deuterium acetone exchange of plasma samples to determine body water enrichment
<ol>
	<li>Reaction
	<ol>
		<li>Prepare 10 standards of deuterium in water, ranging from 0-9% v/v.</li>
		<li>Prepare a 5% v/v acetone/acetonitrile solution allowing for 4 &#181;L per sample, including the standards from step 5.1.1.</li>
		<li>In labelled, safe-lock microcentrifuge tubes, combine 10 &#181;L of each plasma sample or standard, 4 &#181;L of 10 M NaOH, and 4 &#181;L of 5% acetone/acetonitrile. Perform this in triplicate for each sample.</li>
		<li>Mix the samples gently by pipetting. Allow the samples to incubate at room temperature overnight.</li>
	</ol>
	</li>
	<li>Extraction
	<ol>
		<li>After incubation, add 450-550 mg of Na2SO4 to each sample.</li>
		<li>In the fume hood, add 600 &#181;L of CHCl3 to each tube and vigorously vortex for 15 s.</li>
		<li>Centrifuge the samples at 300 x g for 2 min.</li>
		<li>Under a fume hood, transfer triplicate, 80 &#181;L aliquots of the supernatant from each sample into labelled, glass GCMS vials with glass inserts and cap tightly.</li>
	</ol>
	</li>
	<li>GCMS analysis
	<ol>
		<li>Separate samples on a column (30 m, 0.25 mm i.d, Agilent DB-35MS) and analyze on the attached mass spectrometer.
		<ol>
			<li>Inject 1 &#181;L of sample in a split/splitless inlet, with a split ratio of 40:1, a helium flow of 1 mL/min, and an inlet temperature of 270 &#176;C.</li>
			<li>Use the following oven parameters: an initial temperature of 60 &#176;C, increase by 20 &#176;C/min increments to 100 &#176;C, increase by 50 &#176;C/min increments to 220 &#176;C, and then hold at 220 &#176;C for 1 min.</li>
			<li>Use the following mass selective detector (MSD) parameters: electron impact ionization mode at 70 eV with select ion monitoring of 58 and 59 m/z. Use a transfer line at 280 &#176;C, an ion source at 230 &#176;C, and a quadrupole at 150 &#176;C. 
			NOTE: Other low-bleed, bonded, crosslinked, mid-polarity columns may be used, but temperature program will vary.</li>
		</ol>
		</li>
		<li>Set the liquid sampler method to begin with a CHCl3 wash and then a randomized sequence of the samples, including additional CHCl3 wash steps every six samples. 
		NOTE: If acetone is used as a needle wash for the autosampler, replace with CHCl3 or hexane and inject multiple blank CHCl3 samples until any contaminating acetone peak is no longer evident in the chromatogram.</li>
		<li>Using this method, an acetone peak elutes at around 1.25 min. Integrate the data and calculate the fractional abundance of acetone enrichment following steps 6.2.6-6.2.7, with an adjustment to analyze peaks containing an m/z ratio of 58-60 as indicated in Table 1.</li>
		<li>Use the standards to generate a standard curve to determine the percentage of body water enrichment (p) of the test samples, based on the fractional enrichment of acetone.</li>
	</ol>
	</li>
</ol>
8. In vivo de novo lipogenesis calculations
<ol>
	<li>Calculate the fraction of newly synthesized fatty acids (FNS) that are direct products of FAS (i.e., palmitate, odd chain fatty acids, and mmBCFAs) in each specimen with the following equation: 
	<img alt="Equation 3" src="/files/ftp_upload/64219/64219eq3v2.png" style="margin: auto; height: 45px;" /> 
	where ME is the average molar enrichment of a palmitate molecule (step 6.2.9), p is the deuterium enrichment in water from the corresponding plasma sample (step 7.3.4), and N is the number of exchangeable hydrogen atoms on palmitate where a deuterium can be incorporated.</li>
	<li>Determine N using the equation below which was established by Lee et al.31: 
	<img alt="Equation 4" src="/files/ftp_upload/64219/64219eq4v2.png" style="margin: auto; height: 55px;" /></li>
	<li>Determine the molar amount of newly synthesized fatty acids (MNS) by: 
	MNS = FNS x total fatty acid amount (nmol/mg tissue). 
	NOTE: For example, if obtaining a palmitate molar enrichment (ME) of 0.245, a deuterium enrichment in body water (p) of 0.045, and a calculated N number of 22, the fractional synthesis of palmitate is 0.247. If the amount of palmitate present in the tissue is 2 mmol/mg, then the mmol of newly synthesized palmitate is 0.494 mmol/mg (see Supplementary File).</li>
</ol>

The authors have nothing to disclosure.

Understanding the balance and interaction between complex metabolic pathways is an indispensable step toward understanding the biological basis of metabolic related diseases. Here, we show a noninvasive and inexpensive methodology to determine changes in de novo fatty acid synthesis. This method is adapted from previously published methods which developed calculations for estimating de novo synthesis flux from fatty acid deuterium enrichment31 and for using deuterium-acetone exch...

Obesity and associated metabolic diseases are a pandemic that endanger present and future generations1,2. Commonly simplified as the consequence of the imbalance between energy intake and expenditure, the metabolic dysregulation associated with obesity affects a large number of metabolic pathways controlled by environmental and endogenous factors3. However, only a few pathways are routinely tested in animal models of metabolic dysregulation.
As an example, glucose disposal is routinely measured by glucose and insulin tolerance tests, probably due to the simplicity of using portable glucose monitors4. Whole body glucose and lipid oxidation relative rates are also routinely estimated based on the respiratory exchange ratio from indirect calorimetry assays5,6. However, the majority of all other aspects of metabolism are not routinely functionally assessed. This leads to incomplete interpretations of the metabolic status and missed therapeutic options. One of the main such pathways is de novo lipogenesis.
De novo lipogenesis (DNL) is the process by which new fatty acids are generated from precursors. Glucose is considered to be the main precursor contributing to whole-body DNL7, however other precursors, such as acetate, fructose, lactate, and branched chain amino acids, have been shown to be relevant carbon sources in a spatial and condition dependent manner8,9,10,11,12. DNL is a key contributor to metabolic homeostasis and is essential for normal development13. Additionally, alterations in DNL have been associated with cancer14,15 and metabolic16,17,18 and cardiovascular diseases19,20.
The DNL pathway is composed of the core enzymatic components ATP citrate lyase (ACLY), acetyl-CoA carboxylase (ACC1/2), and fatty acid synthase (FAS) which primarily produce palmitate, a 16-carbon saturated fatty acid. However, odd chain and branched chain fatty acids can also be produced at lower rates9. Elongases and desaturases further modify these fatty acids, creating a diverse range of fatty acids species useful for a variety of functions (e.g., long-term energy storage and manipulation of membrane fluidity).
The expression of the DNL enzymatic machinery is controlled by a short number of transcription factors. The most well described to date include the sterol regulatory element binding protein (SREBP) family, carbohydrate response element binding protein (ChREBP), and liver X receptor (LXR)21,22,23,24,25,26. Despite an apparent overlap in their functions, individual regulations based on cell type dominancy and physiological or pathological conditions have been reported21,22,27,28.
Remarkably, a number of inhibitors for selected steps of the DNL pathway have been approved for clinical use or are in the preclinical or clinical stages of development for a number of diseases, including obesity, nonalcoholic fatty liver disease/nonalcoholic steatohepatitis (NAFLD/NASH), and cardiovascular disease29. These efforts highlight the relevance of DNL in health and disease.
In recent years, the employment of methods to quantitatively assess de novo fatty acid synthesis has increased30. The most common method for assessing this is the use of heavy labelled water (D2O), where the heavy labelled hydrogen gets incorporated into acyl chains during synthesis both directly and indirectly, via deuterium exchange with the hydrogens of the DNL substrates NAPDH, acetyl-CoA, and malonyl-CoA. Although this approach is gaining in popularity, there is a lack of publicly available detailed protocols suitable for newcomers in the field. Here, we outline a method for quantitatively assessing the de novo synthesis of products of FAS using D2O and gas chromatography mass spectrometry (GCMS), with calculations previously developed by Lee et al.31. This method measures de novo fatty acid synthesis independently of a carbon source, and it is potentially useful for virtually any tissue, in any mouse model, and under any external perturbation. Here, we focus on the analysis of brown adipose tissue (BAT) due to its high levels of DNL and critical roles in maintaining metabolic homeostasis.

<table><tbody><tr><td>4 mL Glass Vials</td><td>Fisher Scientific</td><td>14-955-334</td><td></td></tr><tr><td>0.2 &amp;micro;m filter</td><td>Olympus Plastic</td><td>25-244</td><td></td></tr><tr><td>26G needeled syringes</td><td>BD</td><td>309597</td><td></td></tr><tr><td>Acetone</td><td>Merck</td><td>34850</td><td></td></tr><tr><td>Acetonitrile</td><td>Merck</td><td>900667</td><td></td></tr><tr><td>Blue GC screw cap with septa</td><td>Agilent</td><td>5190-1599</td><td></td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5424R</td><td></td></tr><tr><td>Chloroform</td><td>Sigma</td><td>366927</td><td></td></tr><tr><td>Deuterium oxide</td><td>Sigma</td><td>151882</td><td></td></tr><tr><td>Di-tert-butyl-4-methylphenol (BHT)&lt;br/&gt; Select FAME Column</td><td>Merck</td><td>B1378</td><td></td></tr><tr><td>Di-tert-butyl-4-methylphenol (BHT)&lt;br/&gt; Select FAME Column</td><td>Agilent</td><td>CP7419</td><td></td></tr><tr><td>EDTA tube</td><td>Sarstedt</td><td>411395105</td><td></td></tr><tr><td>Ethanol</td><td>Merck</td><td>51976</td><td></td></tr><tr><td>Hexadecenoic-d31 Acid</td><td>Larodan</td><td>71-1631</td><td></td></tr><tr><td>Hexane</td><td>Merck</td><td>34859</td><td></td></tr><tr><td>Methanol</td><td>Merck</td><td>34860</td><td></td></tr><tr><td>Microcentrifuge tube</td><td>Olympus Plastic</td><td>24-282</td><td></td></tr><tr><td>Mouse environmental chamber</td><td>Caron</td><td>Caron 7001-33</td><td></td></tr><tr><td>Potasium Chloride</td><td>Fisher Bioreagents</td><td>BP366-500</td><td></td></tr><tr><td>Potasium Phosphate</td><td>MP Biomedicals</td><td>194727</td><td></td></tr><tr><td>SafeLock microcentrifuge tubes</td><td>Eppendorf</td><td>30120086</td><td></td></tr><tr><td>Screw top amber GC vial</td><td>Agilent</td><td>5182-0716</td><td></td></tr><tr><td>Sodium Chloride</td><td>Fisher Bioreagents</td><td>BP358-212</td><td></td></tr><tr><td>Sodium Hydroxide</td><td>Merck</td><td>S5881</td><td></td></tr><tr><td>Sodium Phosphate, dibasic</td><td>Fisher Bioreagents</td><td>BP332-500</td><td></td></tr><tr><td>Sodium Sulfate</td><td>Merck</td><td>239313</td><td></td></tr><tr><td>Sulfuric Acid</td><td>Merck</td><td>258105</td><td></td></tr><tr><td>Vial insert</td><td>Agilent</td><td>5183-2088</td><td></td></tr></tbody></table>

quantitative determination of de novo fatty acid synthesis in brown adipose tissue using deuterium oxide

Fatty acid synthesis is a complex and highly energy demanding metabolic pathway with important functional roles in the control of whole-body metabolic homeostasis and other physiological and pathological processes. Contrary to other key metabolic pathways, such as glucose disposal, fatty acid synthesis is not routinely functionally assessed, leading to incomplete interpretations of metabolic status. In addition, there is a lack of publicly available detailed protocols suitable for newcomers in the field. Here, we describe an inexpensive quantitative method utilizing deuterium oxide and gas chromatography mass spectrometry (GCMS) for the analysis of total fatty acid de novo synthesis in brown adipose tissue in vivo. This method measures the synthesis of the products of fatty acid synthase independently of a carbon source, and it is potentially useful for virtually any tissue, in any mouse model, and under any external perturbation. Details on the sample preparation for GCMS and downstream calculations are provided. We focus on the analysis of brown fat due to its high levels of de novo fatty acid synthesis and critical roles in maintaining metabolic homeostasis.

Based on the D2O dosing described in step 1, we typically find that body water is enriched in the range of 2.5% to 6%, and that a baseline level of deuterium enrichment in body water is rapidly achieved in 1 h and maintained for the duration of the study via 8% enriched drinking water (Figure 1). Continuous steady state body water enrichment is an assumption of the calculations used in step 6, and so we recommend experimental validation of the kinetics of body water enric...

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Quantitative Determination of De Novo Fatty Acid Synthesis in Brown Adipose Tissue Using Deuterium Oxide

Here, we present an inexpensive quantitative method utilizing deuterium oxide and gas chromatography mass spectrometry (GCMS) for the analysis of total fatty acid de novo lipogenesis in brown adipose tissue in vivo.

Quantitative Determination of De Novo Fatty Acid Synthesis in Brown Adipose Tissue Using Deuterium Oxide

Fatty acid synthesis is a complex and highly energy demanding metabolic pathway with important functional roles in the control of whole-body metabolic ...

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966 Views.  University College Dublin. In tegenstelling tot andere metabole routes zoals glucoseverwijdering, wordt de vetzuursynthese niet routinematig beoordeeld, wat leidt tot onvolledige interpretaties van de metabole status. De belangrijkste voordelen van deze methode zijn dat het relatief niet-invasief is en waarschijnlijk een betere weerspiegeling is van de vetzuursyntheseflux dan het gebruik van isotoop-gelabelde glucose, waarbij veranderingen in de stroomopwaartse glucoseverwerking de resultaten kunnen beïnvloeden. Hier ...

Video: Quantitative Determination of De Novo Fatty Acid Synthesis in Brown Adipose Tissue Using Deuterium Oxide

Fatty Acid 13C Isotopologue Profiling Provides Insight into Trophic Carbon Transfer and Lipid Metabolism of Invertebrate Consumers

Fatty acids (FAs) are useful biomarkers in food web ecology because they are typically assimilated as a complete molecule and transferred into consumer tissue with minor or no modification, allowing the dietary routing between different trophic levels. However, the FA trophic marker approach is still hampered by the limited knowledge in lipid metabolism of the soil fauna. This study used entirely labelled palmitic acid (13C16:0, 99 atom%) as a tracer in fatty acid metabolism pathways of two widespread soil Collembola, Protaphorura fimata and Heteromurus nitidus. In order to investigate the fate and metabolic modifications of this precursor, a method of isotopologue profiling is presented, performed by mass spectrometry using single ion monitoring. Moreover, the upstream laboratory feeding experiment is described, as well as the extraction and methylation of dominant lipid fractions (neutral lipids, phospholipids) and the related formula and calculations. Isotopologue profiling does not only yield the overall 13C enrichment in fatty acids derived from the 13C labeled precursor but also produces the pattern of isotopologues exceeding the mass of the parent ion (i.e., the FA molecular ion M+) of each labeled FA by one or more mass units (M+1, M+2, M+3, etc.). This knowledge allows conclusions on the ratio of dietary routing of an entirely consumed FA in comparison to de novo biosynthesis. The isotopologue profiling is suggested as a useful tool for evaluation of fatty acid metabolism in soil animals to disentangle trophic interactions.

Fatty Acid 13C Isotopologue Profiling Provides Insight into Trophic Carbon Transfer and Lipid Metabolism of Invertebrate Consumers

The fatty acid trophic marker approach, i.e., the assimilation of fatty acids as entire molecule and transfer into consumer tissue with no or minor modification, is hampered by knowledge gaps in fatty acid metabolism of small soil invertebrates. Isotopologue profiling is provided as a valuable tool to disentangle trophic interactions.

Fatty acids (FAs) are useful biomarkers in food web ecology because they are typically assimilated as a complete molecule and transferred into consumer ...

Biochemistry

Using Deuterium Oxide as a Non-Invasive, Non-Lethal Tool for Assessing Body Composition and Water Consumption in Mammals

Body condition scoring systems and body condition indices are common techniques used for assessing the health status or fitness of a species. Body condition scoring systems are evaluator dependent and have the potential to be highly subjective. Body condition indices can be confounded by foraging, the effects of body weight, as well as statistical and inferential problems. An alternative to body condition scoring systems and body condition indices is using a stable isotope such as deuterium oxide to determine body composition. The deuterium oxide dilution method is a repeatable, quantitative technique used to estimate body composition in humans, wildlife, and domestic species. Additionally, the deuterium oxide dilution technique can be used to determine the water consumption of an individual animal. Here, we describe the adaption of the deuterium oxide dilution technique for assessing body composition in big brown bats (Eptesicus fuscus) and for assessing water consumption in cats (Felis catis).

This article describes the deuterium oxide dilution technique in two mammals, an insectivore and carnivore, to determine total body water, lean body mass, body fat mass, and water consumption.

Body condition scoring systems and body condition indices are common techniques used for assessing the health status or fitness of a species. Body ...

Environment

Measurement of Fatty Acid &#946;-Oxidation in a Suspension of Freshly Isolated Mouse Hepatocytes

Fatty acid &#946;-oxidation is a key metabolic pathway to meet the energy demands of the liver and provide substrates and cofactors for additional processes, such as ketogenesis and gluconeogenesis, which are essential to maintain whole-body glucose homeostasis and support extra-hepatic organ function in the fasted state. Fatty acid &#946;-oxidation occurs within the mitochondria and peroxisomes and is regulated through multiple mechanisms, including the uptake and activation of fatty acids, enzyme expression levels, and availability of cofactors such as coenzyme A and NAD+. In assays that measure fatty acid &#946;-oxidation in liver homogenates, cell lysis and the common addition of supraphysiological levels of cofactors mask the effects of these regulatory mechanisms. Furthermore, the integrity of the organelles in the homogenates is hard to control and can vary significantly between preparations. The measurement of fatty acid &#946;-oxidation in intact primary hepatocytes overcomes the above pitfalls. This protocol describes a method for the measurement of fatty acid &#946;-oxidation in a suspension of freshly isolated primary mouse hepatocytes incubated with 14C-labeled palmitic acid. By avoiding hours to days of culture, this method has the advantage of better preserving the protein expression levels and metabolic pathway activity of the original liver, including the activation of fatty acid &#946;-oxidation observed in hepatocytes isolated from fasted mice compared to fed mice.

Fatty acid &#946;-oxidation is an essential metabolic pathway responsible for generating energy in many different cell types, including hepatocytes. Here, we describe a method to measure fatty acid &#946;-oxidation in freshly isolated primary hepatocytes using 14C-labeled palmitic acid.

Fatty acid &#946;-oxidation is a key metabolic pathway to meet the energy demands of the liver and provide substrates and cofactors for additional ...

Whole Body and Regional Quantification of Active Human Brown Adipose Tissue Using 18F-FDG PET/CT

In endothermic animals, brown adipose tissue (BAT) is activated to produce heat for defending body temperature in response to cold. BAT&#39;s ability to expend energy has made it a potential target for novel therapies to ameliorate obesity and associated metabolic disorders in humans. Though this tissue has been well studied in small animals, BAT&#39;s thermogenic capacity in humans remains largely unknown due to the difficulties of measuring its volume, activity, and distribution. Identifying and quantifying active human BAT is commonly performed using 18F-Fluorodeoxyglucose (18F-FDG) positron emission tomography and computed tomography (PET/CT) scans following cold-exposure or pharmacological activation. Here we describe a detailed image-analysis approach to quantify total-body human BAT from 18F-FDG PET/CT scans using an open-source software. We demonstrate the drawing of user-specified regions of interest to identify metabolically active adipose tissue while avoiding common non-BAT tissues, to measure BAT volume and activity, and to further characterize its anatomical distribution. Although this rigorous approach is time-consuming, we believe it will ultimately provide a foundation to develop future automated BAT quantification algorithms.

Whole Body and Regional Quantification of Active Human Brown Adipose Tissue Using 18F-FDG PET/CT

Using free, open-source software, we have developed an analytical approach to quantify total and regional brown adipose tissue (BAT) volume and metabolic activity of BAT using 18F-FDG PET/CT.

In endothermic animals, brown adipose tissue (BAT) is activated to produce heat for defending body temperature in response to cold. BAT&#39;s ability to ...

Medicine

Determination of Fatty Acid Oxidation and Lipogenesis in Mouse Primary Hepatocytes

Lipid metabolism in liver is complex. In addition to importing and exporting lipid via lipoproteins, hepatocytes can oxidize lipid via fatty acid oxidation, or alternatively, synthesize new lipid via de novo lipogenesis. The net sum of these pathways is dictated by a number of factors, which in certain disease states leads to fatty liver disease. Excess hepatic lipid accumulation is associated with whole body insulin resistance and coronary heart disease. Tools to study lipid metabolism in hepatocytes are useful to understand the role of hepatic lipid metabolism in certain metabolic disorders.
In the liver, hepatocytes regulate the breakdown and synthesis of fatty acids via &#946;-fatty oxidation and de novo lipogenesis, respectively. Quantifying metabolism in these pathways provides insight into hepatic lipid handling. Unlike in vitro quantification, using primary hepatocytes, making measurements in vivo is technically challenging and resource intensive. Hence, quantifying &#946;-fatty acid oxidation and de novo lipogenesis in cultured mouse hepatocytes provides a straight forward method to assess hepatocyte lipid handling. 
Here we describe a method for the isolation of primary mouse hepatocytes, and we demonstrate quantification of &#946;-fatty acid oxidation and de novo lipogenesis, using radiolabeled substrates.

De novo lipogenesis and &#946;-fatty acid oxidation constitute key metabolic pathways in hepatocyte, pathways that are perturbed in several metabolic disorders, including fatty liver disease. Here we demonstrate isolation of mouse primary hepatocytes and describe quantification of &#946;-fatty acid oxidation and lipogenesis.

Lipid metabolism in liver is complex. In addition to importing and exporting lipid via lipoproteins, hepatocytes can oxidize lipid via fatty acid ...

Methods for the Determination of Rates of Glucose and Fatty Acid Oxidation in the Isolated Working Rat Heart

The mammalian heart is a major consumer of ATP and requires a constant supply of energy substrates for contraction. Not surprisingly, alterations of myocardial metabolism have been linked to the development of contractile dysfunction and heart failure. Therefore, unraveling the link between metabolism and contraction should shed light on some of the mechanisms governing cardiac adaptation or maladaptation in disease states. The isolated working rat heart preparation can be used to follow, simultaneously and in real time, cardiac contractile function and flux of energy providing substrates into oxidative metabolic pathways. The present protocol aims to provide a detailed description of the methods used in the preparation and utilization of buffers for the quantitative measurement of the rates of oxidation for glucose and fatty acids, the main energy providing substrates of the heart. The methods used for sample analysis and data interpretation are also discussed. In brief, the technique is based on the supply of 14C- radiolabeled glucose and a 3H- radiolabeled long-chain fatty acid to an ex vivo beating heart via normothermic crystalloid perfusion. 14CO2 and 3H2O, end byproducts of the enzymatic reactions involved in the utilization of these energy providing substrates, are then quantitatively recovered from the coronary effluent. With knowledge of the specific activity of the radiolabeled substrates used, it is then possible to individually quantitate the flux of glucose and fatty acid in the oxidation pathways. Contractile function of the isolated heart can be determined in parallel with the appropriate recording equipment and directly correlated to metabolic flux values. The technique is extremely useful to study the metabolism/contraction relationship in response to various stress conditions such as alterations in pre and after load and ischemia, a drug or a circulating factor, or following the alteration in the expression of a gene product.

The following protocol describes the preparation and utilization of buffers for the quantitative measurement of rates of glucose and fatty acid oxidation in the isolated working rat heart. The methods used for sample analysis and data interpretation are also discussed.

The mammalian heart is a major consumer of ATP and requires a constant supply of energy substrates for contraction. Not surprisingly, alterations of ...

Using a Combination of Indirect Calorimetry, Infrared Thermography, and Blood Glucose Levels to Measure Brown Adipose Tissue Thermogenesis in Humans

In mammals, brown adipose tissue (BAT) is activated rapidly in response to cold in order to maintain body temperature. Although BAT has been studied greatly in small animals, it is difficult to measure the activity of BAT in humans. Therefore, little is known about the heat-generating capacity and physiological significance of BAT in humans, including the degree to which components of the diet can activate BAT. This is due to the limitations in the currently most used method to assess the activation of BAT-radiolabeled glucose (fluorodeoxyglucose or 18FDG) measured by positron emission tomography-computerized tomography (PET-CT).
This method is usually performed in fasted subjects, as feeding induces glucose uptake by the muscles, which can mask the glucose uptake into the BAT. This paper describes a detailed protocol for quantifying total-body human energy expenditure and substrate utilization from BAT thermogenesis by combining indirect calorimetry, infrared thermography, and blood glucose monitoring in carbohydrate-loaded adult males. To characterize the physiological significance of BAT, measures of the impact of BAT activity on human health are critical. We demonstrate a protocol to achieve this by combining carbohydrate loading and indirect calorimetry with measurements of supraclavicular changes in temperature. This novel approach will help to understand the physiology and pharmacology of BAT thermogenesis in humans.

Here, we present a protocol to quantify the physiological significance of the impact of brown adipose tissue (BAT) activity on human metabolism. This is achieved by combining carbohydrate loading and indirect calorimetry with measurements of supraclavicular changes in temperature. This novel approach can help develop a pharmacological target for BAT thermogenesis in humans.

In mammals, brown adipose tissue (BAT) is activated rapidly in response to cold in order to maintain body temperature. Although BAT has been studied ...

Visualization and Quantification of Brown and Beige Adipose Tissues in Mice using [18F]FDG Micro-PET/MR Imaging

Brown and beige adipocytes are now recognized as potential therapeutic targets for obesity and metabolic syndromes. Non-invasive molecular imaging methods are essential to provide critical insights into these thermogenic adipose depots. Here, the protocol presents a PET/MR imaging-based method to evaluate the activity of brown and beige adipocytes in mouse interscapular brown adipose tissue (iBAT) and inguinal subcutaneous white adipose tissue (iWAT). Visualization and quantification of the thermogenic adipose depots were achieved using [18F]FDG, the non-metabolizable glucose analog, as the radiotracer, when combined with the precise anatomical information provided by MR imaging. The PET/MR imaging was conducted 7 days after cold acclimation and quantitation of [18F]FDG signal in different adipose depots was conducted to assess the relative mobilization of thermogenic adipose tissues. Removal of iBAT substantially increased cold-evoked [18F]FDG uptake in iWAT of the mice.

Visualization and Quantification of Brown and Beige Adipose Tissues in Mice using [18F]FDG Micro-PET/MR Imaging

Functional imaging and quantitation of thermogenic adipose depots in mice using a micro-PET/MR imaging-based approach.

Brown and beige adipocytes are now recognized as potential therapeutic targets for obesity and metabolic syndromes. Non-invasive molecular imaging methods ...

Evaluation of Fatty Acid Oxidation in Cultured Bone Cells

Bone formation by differentiating osteoblasts is expected to require significant energetic input as these specialized cells must synthesize large extracellular matrix proteins that compose bone tissue and then concentrate the ions necessary for its mineralization. Data on the metabolic requirements of bone formation are emerging rapidly. While much remains to be learned, it is expected that derangements in the intermediary metabolism contribute to skeletal disease. Here, a protocol is outlined to assess the capacity of osteoblastic cells to oxidize 14C-labeled fatty acids to 14CO2 and acid-soluble metabolites. Fatty acids represent a rich-energy reserve that can be taken up from the circulation after feeding or after their liberation from adipose tissue stores. The assay, performed in T-25 tissue culture flasks, is helpful for the study of gene gain or loss-of-function on fatty acid utilization and the effect of anabolic signals in the form of growth factors or morphogens necessary for the maintenance of bone mass. Details on the ability to adapt the protocol to assess the oxidation of glucose or amino acids like glutamine are also provided.

The present protocol describes an assay to assess the capacity for fatty acid oxidation in cultures of primary bone cells or relevant cell lines.

Bone formation by differentiating osteoblasts is expected to require significant energetic input as these specialized cells must synthesize large ...

Differentiation and Imaging of Brown Adipocytes from the Stromal Vascular Fraction of Interscapular Adipose Tissue from Newborn Mice

Brown adipose tissue (BAT) is only present in mammals and has a thermogenic function. Brown adipocytes are characterized by a multilocular cytoplasm with multiple lipid droplets, a central nucleus, a high mitochondrial content, and the expression of uncoupling protein 1 (UCP1). BAT has been proposed as a potential therapeutic target for obesity and its associated metabolic disorders due to its ability to dissipate metabolic energy as heat. To investigate BAT function and regulation, brown adipocyte culturing is indispensable. The present protocol optimizes tissue processing and cell differentiation for culturing brown adipocytes from newborn mice. Additionally, procedures for the imaging of differentiated adipocytes with both confocal immunofluorescence and transmission electron microscopy are shown. In the brown adipocytes differentiated with the techniques described herein, the major defining features of classical BAT are preserved, including high UCP1 levels, increased mitochondrial mass, and very close physical contact between the lipid droplets and mitochondria, making this method a valuable tool for BAT studies.

Preadipocytes are isolated from the stromal vascular fraction of interscapular brown adipose tissue from newborn mice and differentiated into cells that accumulate lipid droplets, express molecular markers, and show the mitochondrial morphology of mature brown adipocytes. These cells are further analyzed by immunofluorescence and transmission electron microscopy.

Brown adipose tissue (BAT) is only present in mammals and has a thermogenic function. Brown adipocytes are characterized by a multilocular cytoplasm with ...

Quantitative Determination of De Novo Fatty Acid Synthesis in Brown Adipose Tissue Using Deuterium Oxide

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